Phosphoinositide Metabolism and Signaling During Late Phagosome Maturation and Phagolysosome Resolution

by

Roni Levin Konigsberg

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

Department of Biochemistry University of Toronto

© Copyright by Roni Levin Konigsberg 2018

ii

Phosphoinositide Metabolism and Signaling During Late

Phagosome Maturation and Phagolysosome Resolution

Roni Levin Konigsberg

Doctor of Philosophy

Department of Biochemistry University of Toronto

2018

Abstract

Phagocytosis, the regulated uptake of particulate matter by cells, is essential to immunity and

homeostasis. Phagocytes are tasked with the immune surveillance that prevents infection by

invading pathogens. Additionally, these cells clear accumulated apoptotic debris that results

from the daily turnover of billions of cells. In order to fulfill these paramount functions,

phagocytes recognize targets and internalize them into membrane-bound compartments termed

phagosomes. Following internalization, cells degrade the phagosomal contents and dispose of

them resuming the immune response. Phagocytosis is therefore divided into three stages:

formation, maturation and resolution. Formation refers to the recognition and engulfment of

prey. The transformation that enables degradation of the target is termed maturation. Finally,

the elimination and often re-utilization of degraded contents is known as resolution. Most of the

phenomena that govern each of these stages are orchestrated by acute signaling events.

Specialized signaling phospholipids known as phosphoinositides mediate the recruitment of

effector proteins to phagosomes. While the roles of diverse phosphoinositides during formation

and the early maturation have been studied and partially elucidated, little is known about them

during the late maturation and resolution stages. The work described in this dissertation was

conducted with the aim of characterizing phosphatidylinositol 4-phosphate (PtdIns4P)—a

phosphoinositide present in the plasma membrane, Golgi apparatus and late compartments—

iii

during phagocytosis. Specifically, I focused on elucidating the dynamics, metabolism and

functions of PtdIns4P during each stage of phagocytosis. Chapter I, consists of relevant

background on the cell biology of phagocytosis and the involvement of phosphoinositides during

the process. The aims and hypothesis of my work are described in Chapter 2. Chapter 3

describes the methodology used throughout this work. In Chapter 4, I detail PtdIns4P dynamics

and metabolic mechanisms during phagosome formation and maturation. Additionally, I define

the functional implications of PtdIns4P in maturing phagosomes. In Chapter 5 I detail how the

endoplasmic reticulum regulates both phagosomal PtdIns4P levels and recruitment of its

effectors to the compartment. Finally, I explore the relevance of these events during resolution.

This work establishes the importance of PtdIns4P during phagocytosis emphasizing the

necessity of an exquisite coordination of lipid metabolism during the process.

iv

Acknowledgments I grew up playing basketball. As a player, I considered myself a “pass-first” point guard. My

favorite play was always—and still is—to give a pass to a teammate leading to a basket. This

play is accurately termed an “assist”. Ironically, for one of the biggest achievements of my life, I

am the one who has been assisted—vastly. This journey would not have been possible without

the constant help and massive support of many people. Words will never suffice to genuinely

express my gratitude towards each one of them. Below is my attempt.

First, I want to thank my wife Rossie, my best friend. For your patience that has allowed me to

follow my dream; for your support that keeps me going everyday; and more than anything for

your love. You’re my motor, my direction and my balance. You joined my adventures, now we

have an amazing life together and an exciting future together. I cannot wait for our many

adventures to come.

I want to thank my parents Elisa and Sergio; I owe you everything I am and everything I’ve

done. Any achievement of mine will always be yours. My main objective in life is to reflect both

of you in everything I do, I truly admire you; you will forever be my role models. Words do not

exist to thank you for what you have done for me. Thank you for showing me how to live and for

your never-ending support.

I want to thank my brothers, Gabriel and Yair; you are my inspiration, for a long time, but these

days more than ever. Seeing you achieve your goals and fulfill your dreams fills me with pride.

While taking different professional paths, we will always be connected by the same values and

our approach to life. I also want to welcome Mariana to our family.

I want to thank my grandmother Fanny and my Bobe Clara, for always being there. For all your

love and support throughout my entire life. Seeing you and talking to you is always special.

There are far too many things that make both of you unique and admirable.

I not only want to thank, but I also want to dedicate this thesis to my grandfather Alfredo, to my

Zeide Sommer and to my father-in-law Salo. Your memories are a true blessing to me. Getting

to know you filled my heart; I wish all of you were here. Alfredo, thank you for your guidance,

wisdom and endless love, I never stopped admiring you. Zeide, thank you for teaching me the

true meaning of perseverance and showing how to cope with real adversity, you will always be

my motivation. Salo, thank you for sharing you joy for life with me and everyone. For enjoying

v

every moment and always laughing out loudly. Thank you for trusting me to share my life with

your daughter’s.

Thank you Sergio; saying that more than a supervisor, you have been a wonderful mentor and

an outstanding teacher is an understatement. Not for a single day during this journey I took your

guidance for granted. Since day one I understood how privileged I am to be your trainee. I have

learned from you on a daily basis, yet I know that I would never stop learning something new

from El Profe. It is not often that you find a person with so many qualities in all fronts. It seems

impossible to imagine a better mentor. From how to write to fundamental physiology and

biochemistry, from how to speak during presentations to advanced microscopy techniques, from

having an open door on a daily basis at the lab to shooting hoops during a conference, I will

forever cherish each moment.

Thank you to my friends, Lanny, Jessi, Charlie, Charlie, Jeremy, Jason, Alec, Mitchell, Sarah,

Julie and Ben. I couldn’t have been luckier to have such a wonderful group of brothers and

sisters. You have been a true family to us, thank you for ‘adopting’ us, and for such amazing

times together. I will miss you everyday, but I’m happy knowing that our friendships will last a

lifetime. You are the best friends anyone could ever dream of.

I want to thank Greg; while not being my supervisor ‘on paper’, you certainly acted like a

mentor—a great one—throughout my Ph. D. The only comparable thing to your vast knowledge

is your kindness. Thank you for the sacrifices you made in order to be there for weekly

meetings, technical support, ‘brain-picking’, even ‘babysitting’ sometimes. I really enjoyed

talking science and sports regularly. I am also lucky and honored you included me as part of

your lab. Watching your lab grow over these years has been special.

I would like to thank Peter Kim, as my committee member. I’m grateful for your constant advice

on both, science and life. I always enjoy our conversations whenever we run into each other.

You have certainly helped me through my doctoral studies and for my future as a postdoc.

Thank you Dr. Roberto Zoncu, for agreeing to be my external examiner. I am truly honored that

you took the time to do this, including writing an appraisal for this work.

I want to thank all of my collaborators, whose crucial contributions to this work are detailed

below. It was a tremendously positive experience and I am fortunate you are part of this work. I

am especially grateful to Gerry and Tamas; this work would not have possible without your

willingness to help, your expertise, as well as your knowledge and advice.

vi

I want to thank Sharon and Takashi, who were like family to me since the beginning. I am happy

to have crossed paths with you and for our beautiful friendship. Thank you for making me feel at

home and as a part of your family anytime I was with you. Similarly, I would like to thank

Fernando and Paul (and Melody). While both of you joined the lab as I was closer to finishing,

I’m grateful that I got to meet you and for our special friendship.

To all my friends in the Grinstein lab—past and current members. Specially, Stella (my grad

school buddy), Sivakami (also for critically reading and editing sections of this dissertation),

Pedro, Ziv, Johannes, Glenn, Johnathan, Rich, Phil, Cat, Dasha, JP. Thank you for creating

such a lovely environment on a daily basis. It has been a true pleasure to work by your side

these years. Thanks for the memories, the feedback and constant help. More than anything

thanks for the laughs ‘on and off’ the lab. I truly cherish these friendships and I hope they go on.

Additionally, I want to thank the wonderful friends I met from other labs: Mariana, Mauricio,

Javier and Kitty (who became like family to us), Tara, Nick, Rafaela, Cheryl, Scott, Wael, Ren

and Rachel.

I also want to thank Rashna, Sheryl and Carrie. For always being willing to help—very

efficiently—with a big smile. Thanks to you these years went as smooth as they could. I always

enjoyed chatting with each one of you while you were helping me with something.

I want to thank Michael Bassik. For trusting me and accepting me into your lab. I am excited to

join your team, looking forward to doing science together and learning from you.

Finally, I would like to thank CONACYT and the Connaught International Scholarship for

doctoral students for funding my Ph. D. training during the past five years.

vii

Table of Contents

Acknowledgments ............................................................................................................ iv

Table of Contents ............................................................................................................ vii

List of Abbreviations ........................................................................................................ xii

List of Figures................................................................................................................ xxii

Contributions ................................................................................................................. xxv

Chapter 1 Introduction ..................................................................................................... 1

General introduction: Phosphoinositide dynamics, metabolism and signaling 1during the life cycle of phagosomes ............................................................................ 1

1.1 Summary ......................................................................................................... 1

1.2 Introduction ..................................................................................................... 2

1.2.1 Stages of phagocytosis .......................................................................... 3

1.2.2 Phosphoinositides in phagocytosis ........................................................ 3

1.3 Stage 1: Phagosome formation ...................................................................... 4

1.3.1 Engaging the target ................................................................................ 4

1.3.2 Transducing the signal ........................................................................... 9

1.3.3 Cytoskeletal rearrangements ............................................................... 13

1.3.4 Sealing the phagosomal membrane ..................................................... 15

1.3.5 Terminating the signal .......................................................................... 16

1.4 Stage 1: Phagosome maturation .................................................................. 17

1.4.1 The early phagosome ........................................................................... 18

1.4.2 Early to late phagosome transition ....................................................... 22

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1.4.3 Phagolysosome biogenesis .................................................................. 26

1.5 Stage 3: Phagosome resolution .................................................................... 28

1.5.1 Disposal of nucleic acids ...................................................................... 29

1.5.2 Protein and amino acid resolution ............................................................ 31

1.5.3 Lipid processing ................................................................................... 32

1.6 Phosphoinositides in phagocytosis ............................................................... 36

1.6.1 Phosphoinositides during phagosome formation ................................. 37

1.6.2 Phosphoinositides during phagosome maturation ............................... 53

1.6.3 Phosphoinositides during phagosome resolution ................................. 64

1.7 Rationale ....................................................................................................... 65

Chapter 2 Hypothesis and Aims .................................................................................... 67

Thesis hypothesis and aims ...................................................................................... 67 2

2.1 Hypothesis .................................................................................................... 67

2.2 Aims .............................................................................................................. 67

Chapter 3 Materials and Methods .................................................................................. 69

General methods ....................................................................................................... 69 3

3.1 Introduction ................................................................................................... 69

3.2 Reagents ....................................................................................................... 70

3.3 Cell culture .................................................................................................... 71

3.4 Primary cell isolation and differentiation ....................................................... 72

3.5 Antibodies ..................................................................................................... 72

3.6 Plasmids ....................................................................................................... 73

3.7 Transient transfections .................................................................................. 74

3.8 Particle opsonization ..................................................................................... 74

ix

3.9 Phagocytosis assay ...................................................................................... 75

3.10 Gene silencing .............................................................................................. 75

3.11 Quantitative RT-PCR .................................................................................... 76

3.12 Generation of CRISPR KO cell lines ............................................................. 77

3.13 Gene editing measurements by Sanger sequencing .................................... 77

3.14 Detection of acidification and lysosomal labeling .......................................... 77

3.15 Confocal microscopy ..................................................................................... 78

3.16 Lattice light-sheet microscopy ....................................................................... 78

3.17 Transmission electron microscopy ................................................................ 79

3.18 Image processing .......................................................................................... 79

3.19 Protein purification ........................................................................................ 80

3.20 Protein-lipid overlay assay ............................................................................ 80

Chapter 4 Multiphasic Dynamics of Phosphatidylinositol 4-phosphate During Phagocytosis ............................................................................................................. 82

Multiphasic dynamics of phosphatidylinositol 4-phosphate during phagocytosis ...... 82 4

4.1 Abstract ......................................................................................................... 82

4.2 Introduction ................................................................................................... 83

4.3 Results .......................................................................................................... 85

4.3.1 Detection of PtdIns4P in macrophages ................................................ 85

4.3.2 PtdIns4P dynamics during phagosome formation and maturation ....... 88

4.3.3 Disappearance of PtdIns4P from the phagosome ................................ 93

4.3.4 PtdIns4P reappearance in maturing phagosomes ............................... 99

4.3.5 PtdIns4P is required for completion of phagosome maturation .......... 103

4.4 Discussion ................................................................................................... 108

x

Chapter 5 Phosphatidylinositol 4-phosphate Regulation by Endoplasmic Reticulum – Phagolysosome Contact Sites Directs Phagosome Resolution .............................. 112

Phosphatidylinositol 4-phosphate regulation by endoplasmic reticulum – 5phagolysosome contact sites directs phagosome resolution ................................... 112

5.1 Abstract ....................................................................................................... 112

5.2 Introduction ................................................................................................. 113

5.3 Results ........................................................................................................ 115

5.3.1 Phagosome resolution ........................................................................ 115

5.3.2 PtdIns4P dynamics during the early stages of phagosome

resolution ........................................................................................................... 117

5.3.3 PtdIns4P degradation mechanism ..................................................... 119

5.3.4 ORP1L-dependent phagolysosome-to-ER PtdIns4P transport .......... 125

5.3.5 Functional implications of PtdIns4P in the phagolysosome ............... 131

5.4 Discussion ................................................................................................... 139

Chapter 6 General Discussion ..................................................................................... 144

Summary of findings, future directions and concluding remarks ............................. 144 6

6.1 PtdIns4P dynamics during phagocytosis .................................................... 144

6.1.1 Summary of findings ........................................................................... 144

6.1.2 Future directions ................................................................................. 145

6.2 PtdIns4P metabolism mechanisms during phagocytosis ............................ 145

6.2.1 Summary of findings and future directions ......................................... 145

6.3 Functional implications of PtdIns4P during phagocytosis ........................... 149

6.3.1 Summary of findings and future directions ......................................... 149

6.4 Concluding remarks .................................................................................... 151

References ................................................................................................................... 152

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Appendix I .................................................................................................................... 181

Copyright Acknowledgements ..................................................................................... 182

xii

List of Abbreviations

ABC ATP-binding cassette

Abt1 Activator of basal transcription

Akt Protein kinase B

AP-1 Adaptor protein complex 1

AP-3 Adaptor protein complex 3

APPL1 Adaptor protein, phosphotyrosine interacting with PH domain and

leucine zipper 1

Arf ADP ribosylation factor

Arl ADP ribosylation factor like

Arp2/3 Actin-related protein 2 and 3

BAR Bin/Amphiphysin/Rvs

Bcl10 B-cell lymphoma/leukemia 10

bec-1 Beclin homolog

BFP Blue fluorescent protein

Bin2 Bridging integrator 2

BSA Bovine serum albumin

Ca2+ Calcium cation

CaSR Calcium sensing receptor

CCP Clathrin coated pit

xiii

CD36 Cluster of differentiation 36

Cdc42 Cell division control protein 42 homolog

CDKN1A Cycling dependent kinase inhibitor

cDNA Chromosomal DNA

CFP Cyan fluorescent protein

CO2 Carbon dioxide

CORVET Class C core vacuole/endosome tethering

CRIB Cdc42- and Rac-interactive binding

CRISPR Clustered regularly interspaced short palindromic repeats

Csk C-terminal Src kinase

CX3CR1 CX3C chemokine receptor 1

DAG Diacylglycerol

DMEM Dulbecco's modified eagle media

DNA Deoxyribonucleic acid

DOCK1 Dedicator of cytokinesis protein 1

EEA1 Early endosome antigen 1

ENT-3 Equilibrative nucleoside transporter 3

ENTH Epsin N-terminal homolog

EPEC Enteropathogenic Escherichia coli

ER Endoplasmic reticulum

xiv

ERM Ezrin/Radixin/Moesin

ESCRT Endosomal sorting complexes required for transport

F-BAR Fes/CIP4 homology BAR

FBP17 Formin-binding protein 17

FcγR Fc gamma receptor

FERM 4.1 protein ERM

FFAT Two phenylalanines in an acidic tract

FIG4 Factor-induced gene 4

FKBP F506 binding protein

FRB FKBP-rapamycin-binding

FYVE Fab1 YOTB Vac1 EEA1

Gab2 GRB2-associated-binding protein

GAPDH Glyceraldehyde 3-phosphate dehydrogenase

GAPex-5 GTPase activating protein and Vps9 activating domains

GDI GDP dissociation inhibitor

GEF Guanine nucleotide exchange factor

GFP Green fluorescent protein

GM1 Monosialic ganglioside 1

GM2 Monosialic ganglioside 2

GM2-AP GM2 activator protein

xv

GM3 Monosialic ganglioside 3

GPCR G protein-coupled receptor

Grb2 Growth factor receptor-bound protein 2

GST Glutathione S-transferase

GTPase Guanosine triphophatase

H2O2 Hydrogen peroxide

HBSS Hank's balanced salt solution

HI-FBS Heat inactivated fetal bovine serum

HOCl Hypochlorus acid

HOPS Homotypic fusion and sorting

HRBC Human red blood cells

HRP Horseradish peroxidase

Hs1 Hematopoietic lineage cell-specific protein 1

Hv1 Voltage-gated proton channel

I-BAR Inverse BAR

IgG Immunoglobulin G

ILV Intraluminal vesicle

INPP5 Inositol polyphosphate 5-phosphatase

Ins(1,4,5)P3 Inositol 1,4,5-trisphosphate

ITAM Immunoreceptor tyrosine-based activation motif

xvi

ITIM Immunoreceptor tyrosine-based inhibition motif

KO Knockout

LAMP Lysosomal-associated membrane protein

LAT Linker for activation of T-cells

LLSM Lattice light-sheet microscopy

LPLA2 Lysosomal phospholipase A2

LPS Lipopolysaccharide

M6PR Mannose 6-phosphate receptor

ManLAM Mannose-capped lipoarabinomannan

MARCO Macrophage receptor with collagenous structure

mCh Monomeric cherry

MCSF Macrophage colony stimulating factor

MerTK Tyrosine-protein kinase MER precursor

MPO Myeloperoxidase

mRFP Monomeric red fluorescent protein

MTM Myotubularin

MTOC Microtubule organizing center

mTORC Mammalian target of rapamycin complex

N-BAR N-terminal amphipathic helix-containing BAR

N-WASP Neural-WASP

xvii

NADPH Reduced nicotinamide adenosine dinucleotide phosphate

Nap1 Nck-associated protein-1

NOX NADPH oxidase

NOX2 NADPH oxidase 2

NPC-1 Niemann-Pick C1 protein

NPC-2 Niemann-Pick C2 protein

NPF Nucleation-promoting factor

OCRL Oculocerebrorenal syndrome of Lowe

ORD OSBP-related domain

ORP1L OSBP-related protein 1L

ORP1S OSBP-related protein IS

OSBP Oxysterol binding protein

Osbpl1 OSBP-like protein 1L

P/S Penicillin streptomycin

P4M PI4P binding of SidM

PBS Phosphate-buffered saline

PBS-T PBS-tween 20

PCR Polymerase chain reaction

PFA Paraformaldehyde

pH Potential hydrogen

xviii

PH Pleckstrin homology

PI3K Phosphatidylinositol 3 kinase

PI4K Phosphatidylinositol 4 kinase

PIKFYVE Phosphatidylinositol 3-phosphate 5-kinase

PIP5K Phosphatidylinositol phosphate 5 kinase

PIPK Phosphatidylinositol phosphate kinase

PKC Protein kinase C

PLC Phospholipase C

PLD Phospholipase D

PM Plasma membrane

PtdCho Phosphatidylcholine

PtdIns Phosphatidylinositol

PtdIns(3,4,5)P3 Phosphatidylinositol 3,4,5-trisphosphate

PtdIns(3,5)P2 Phosphatidylinositol 3,5-bisphosphate

PtdIns(4,5)P2 Phosphatidylinositol 4,5-bisphosphate

PtdIns3P Phosphatidylinositol 3-phosphate

PtdIns4P Phosphatidylinositol 4-phosphate

PtdIns5P Phosphatidylinositol 5-phosphate

PtdOH Phosphatidic acid

PTEN Phosphatase and tensin homolog

xix

PX Phox homology

qPCR Quantitative PCR

RCP Rab coupling protein

RILP Rab7-interacting lysosomal protein

RNA Ribonucleic acid

ROS Reactive oxygen species

RPMI Roswell Park Memorial Institute medium

RT Room temperature

RT-PCR Reverse transcription PCR

SAC Suppressor of actin

SapM Secreted acid phosphatase of Mycobacterium tuberculosis

SCV Salmonella containing vacuole

SEM Standard error of mean

SFK Src family kinases

sgRNA Single guide RNA

SH2 Src homology 2

SH3 Src homology 3

SH3BP2 SH3 domain binding protein 2

SHIP SH2 domain-containing inositol phosphatase

SHP-1 Src homology region 2 domain-containing phosphatase-1

xx

SHP-2 Src homology region 2 domain-containing phosphatase-2

siRNA Short interference RNA

SLC Solute carrier

SNARE

Soluble N-ethylmaleimide sensitive factor attachment protein

receptor

SNX Sorting nexin

SOD Superoxide dismutase

spp Species

SR-B2 Scavenger receptor B2

SRBC Sheep red blood cell

STIM1 Stromal interaction molecule 1

Syk Spleen tyrosine kinase

T3SS Type III secretion system

TAPP1 Tandem pleckstrin homology domain-containing protein 1

TEM Transmission electron microscopy

TGN Trans Golgi network

TIAM1 T-cell lymphoma invasion and metastasis-inducing protein

TIM4 T-cell immunoglobulin- and mucin- domain containing protein 4

TLR Toll-like receptor

TMR Tetramethylrhodamine

xxi

TTC7 Tetratricopeptide repeat domain 7A

V-ATPase Vacuolar ATPase

Vamp Vesicle-associated membrane protein

VAPA Vamp-associated protein A

VAPB Vamp-associated protein B

Vps Vacuolar protein sorting

WASH Wiskott-Aldrich syndrome protein and scar homolog

WASP Wiskott-Aldrich syndrome protein

WAVE WASP-family verprolin homolog

xxii

List of Figures

Chapter 1 - General introduction: phosphoinositide dynamics, metabolism and

signaling during the life cycle of phagosomes ............................................................. 1

Figure 1.1. Phagosome formation ............................................................................... 7

Figure 1.2. Early maturation: from the new phagosome to the early phagosome .... 19

Figure 1.3. Early to late phagosome transition and phagolysosome biogenesis ...... 23

Figure 1.4. Differences in phagosomes between phagocytes .................................. 27

Figure 1.5. Phagosome resolution ............................................................................ 30

Figure 1.6. Distribution of PtdIns(4,5)P2 during phagocytosis ................................... 40

Figure 1.7. Functional implications of PtdIns(4,5)P2 metabolism for phagocytosis ................................................................................................................................... 42

Figure 1.8. Distribution of PtdIns(3,4,5)P3 during phagocytosis ................................ 50

Figure 1.9. Functional implications of PtdIns(3,4,5)P3 metabolism for phagocytosis .............................................................................................................. 52

Figure 1.10. Distribution of PtdIns3P during phagocytosis ....................................... 55

Figure 1.11. Functional implications of PtdIns3P metabolism for phagocytosis ....... 58

Chapter 4 - Multiphasic dynamics of phosphatidylinositol 4-phosphate during

phagocytosis .............................................................................................................. 82

Figure 4.1. PtdIns4P undergoes triphasic changes during phagocytosis ................. 86

Figure 4.2. 2xP4M expression does not affect Golgi morphology ............................ 88

Figure 4.3. Comparison of the changes in PtdIns4P content to those of PtdIns(4,5)P2 and PtdIns3P ...................................................................................... 90

Figure 4.4. Phosphoinositide metabolism during the early stages of phagocytosis ................................................................................................................................... 92

Figure 4.5. Late phagosomes and phagolysosomes contain PtdIns4P .................... 94

xxiii

Figure 4.6. Sac2 recruitment to phagosomes and PtdIns4P degradation ................ 96

Figure 4.7. Assessment of the role of PLC in PtdIns4P disappearance ................... 98

Figure 4.8. PI4K2A recruitment and generation of PtdIns4P in maturing phagosomes ............................................................................................................ 100

Figure 4.9. PtdIns4P dynamics during phagocytosis in COS-1-FcγRIIa cells ........ 102

Figure 4.10. Phagosome acidification is impaired when PI4K2A is silenced .......... 104

Figure 4.11. Cresyl violet co-localizes with Rab7 but not with Rab5 ...................... 105

Figure 4.12. RILP acquisition is impaired in PtdIns4P-depleted phagosomes ....... 107

Figure 4.13. Diagrammatic representation of the changes undergone by phosphoinositides during phagosome formation and maturation ............................ 109

Chapter 5 – Phosphatidylinositol 4-phosphate regulation by endoplasmic reticulum – phagolysosome contact sites directs phagosome resolution ................ 112

Figure 5.1. Phagosomal resolution is characterized by tubular fission events ....... 116

Figure 5.2. Dispersed compartments during phagosome resolution are of phagosomal origin ................................................................................................... 118

Figure 5.3. Dynamics and distribution of PtdIns4P during initiation of phagosome resolution ................................................................................................................. 120

Figure 5.4. PI4K2A remains in the phagosomal membrane during maturation and initiation of phagosome resolution ........................................................................... 121

Figure 5.5. ORP1L accumulates in phagosomes during late maturation ................ 123

Figure 5.6. Monocytes and macrophages express low levels of ORP1S ............... 124

Figure 5.7. ORP1L expression accelerates late PtdIns4P disappearance in maturing phagosomes ............................................................................................. 126

Figure 5.8. PtdIns4P and ORP1L localize to mutually exclusive phagosomal microdomains .......................................................................................................... 127

Figure 5.9. ORP1L KO impairs late PtdIns4P disappearance from maturing phagosomes ............................................................................................................ 128

Figure 5.10. Additional ORP1L KO clones impair PtdIns4P disappearance ........... 130

Figure 5.11. ORP1L binds and transports PtdIns4P to the ER ............................... 132

xxiv

Figure 5.12. ORP1L – VAPA/B interactions mediate phagosome – ER contacts ................................................................................................................................. 134

Figure 5.13. PtdIns4P directly binds SKIP and recruits it to the phagosome .......... 135

Figure 5.14. PtdIns4P stabilizes the Arl8b – SKIP interaction in the phagosomal membrane ................................................................................................................ 137

Figure 5.15. ORP1L mediates phagosome resolution by regulating phagosomal PtdIns4P .................................................................................................................. 138

Figure 5.16. PtdIns(4,5)P2 is not present in phagosomes during PtdIns4P-positive tubulation ................................................................................................................. 140

Figure 5.17. Working model .................................................................................... 143

xxv

Contributions

I was responsible for the execution of the vast majority of the experiments described in this

dissertation. These experiments were conceived and designed by Dr. Sergio Grinstein, Dr.

Gregory Fairn and I. Dr. Grinstein and Dr. Fairn assisted in the interpretation and analysis the

resulting data of all these experiments. Additionally, they played essential roles thoroughly

reviewing and editing the manuscripts that resulted from this work.

Chapter 1. General introduction: phosphoinositide dynamics metabolism and signaling during

the life cycle of the phagosome.

Dr. Johnathan Canton and Dr. Daniel Schlam reviewed and collated primary literature,

wrote parts of independent manuscripts and assisted with the generation of figures.

Chapter 3. Materials and methods.

Dr. Fernando Montaño-Rendón reviewed and collated primary literature and wrote part

of the manuscript.

Chapter 4. Multiphasic dynamics of phosphatidylinositol 4-phosphate during phagocytosis

Dr. Gerald Hammond and Dr. Tamas Balla developed the PtdIns4P sensing probes,

served as scientific consultants and proposed critical experiments. Dr. Hammond, Dr.

Balla and Dr. Pietro de Camilli shared reagents, reviewed and edited the manuscript that

resulted from the work.

Chapter 5. Multiphasic dynamics of phosphatidylinositol 4-phosphate during phagocytosis

Dr. Fernando Montaño-Rendón assisted with cell culture and performed crucial

experimental repeats. Dr. Tal Keren-Kaplan purified SKIP and performed lipid-binding

assays. Braden Ego and Dr. Michael Bassik generated CRISPR cell lines and their

validation. Dr. John Heddleston and Satya Khuon trained and assisted me in lattice light-

sheet microscopy. Jess Diciccio performed electron microscopy experiments and

quantified phagosome resolution stages. Dr. William Trimble, Dr. Juan Bonifacino, Dr.

Keren-Kaplan and Braeden Ego reviewed and edited the manuscript.

1

Chapter 1 Introduction

This chapter includes material from the following:

- Roni Levin, Sergio Grinstein and Johnathan Canton. “The life cycle of phagosomes:

formation, maturation and resolution.” Immunological Reviews – Neutrophils. August 2016.

- Roni Levin, Sergio Grinstein and Daniel Schlam. “Phosphoinositides in Phagocytosis and

Macropinocytosis.” Biochimica et Biophysica Acta – Molecular and Cell Biology of Lipids.

September 2014.

General introduction: Phosphoinositide dynamics, 1metabolism and signaling during the life cycle of phagosomes

1.1 Summary

Professional phagocytes provide immunoprotection and aid in the maintenance of tissue

homeostasis. They perform these tasks by recognizing, engulfing and eliminating pathogens

and endogenous cell debris. This process—termed phagocytosis—is essential for tissue

homeostasis and is also an early, critical component of the innate immune response.

Phagocytosis can be conceptually divided into three stages: phagosome formation, maturation

and resolution. Here, I examine the paramount roles played by phosphoinositides during each

one of these stages. I analyze accumulating literature describing the molecular mechanisms

whereby phosphoinositides translate environmental cues into the complex, sophisticated

responses that underlie the phagocytic response. After describing the subcellular distribution of

phosphoinositides and detailing their mechanisms of synthesis and degradation, I emphasize

the physiological implications of acute changes of their dynamics and metabolism for regulation

of phagocytosis. In addition, I exemplify virulence strategies involving modulation of host cell

phosphoinositide signaling that are employed by bacteria to undermine immunity.

2

1.2 Introduction

Phagocytosis is defined as the regulated uptake of large particles (>0.5 µm in diameter) into

cytosolic, membrane-bound vacuoles called phagosomes. It is an evolutionarily ancient

process used by unicellular organisms such as Dictyostelium spp. as a means of nutrient

acquisition (Bloomfield et al., 2015). In metazoans, dedicated cells termed phagocytes also

utilize phagocytosis for the uptake and recycling of nutrients from effete cells. The functionality

of this process, however, has expanded to include a role in the clearance of a staggering array

of debris in order to maintain homeostasis. Notable examples include the extrusion of

erythroblast nuclei directly onto the surface of macrophages for immediate uptake (Manwani

and Bieker, 2008), the clearance of outer segments shed by photoreceptor cells engulfed by

nearby retinal pigment epithelial cells (Guo et al., 2015) and the removal of senescent red blood

cells from circulation by macrophages in the spleen (Gottlieb et al., 2012).

Phagocytosis in metazoans, though, is not only involved in ‘housekeeping’; it is also

fundamentally important to the maintenance of immune homeostasis. Indeed, the prototypical

phagocytes in mammals are cells of the innate immune system—neutrophils, macrophages and

dendritic cells. During infections, neutrophils are most often the first cells on the scene,

surveying for invading microorganisms. Their capacity to generate and deliver microbicidal

compounds into the phagosome underlies the elimination of potential pathogens. Macrophages

can also clear pathogens at sites of infection, but serve the additional function of cleaning up the

collateral damage caused by the exuberant microbicidal response of neutrophils (Farrera and

Fadeel, 2013). Dendritic cells, and some types of macrophages, adeptly extract antigen from

material digested in phagosomes for presentation to lymphocytes, thereby eliciting

immunological memory (Mantegazza et al., 2014; Alloatti et al., 2015). Thus, phagocytosis is a

central process to both development and homeostasis.

3

1.2.1 Stages of phagocytosis

Phagocytosis is initiated by engagement of surface receptors that recognize ligands exposed by

the target particle. Upon lateral clustering, phagocytic receptors initiate signaling cascades that

culminate in the extension of pseudopod-like structures. These membrane protrusions surround

the target and seal at their distal tips, thereby generating a nascent phagosome. Neutrophils,

macrophages and dendritic cells express distinct but overlapping sets of phagocytic receptors.

Possessing unique receptor repertoires enables individual phagocyte types to recognize certain

targets preferentially. The newly formed phagosome then undergoes a series of fusion and

fission interactions with cytosolic organelles, resulting in extensive remodeling of both its limiting

membrane and luminal contents. This remodeling sequence is collectively referred to as

phagosome maturation; of note, maturation can vary drastically among phagocytes. Finally, the

phagocyte must digest and either recycle or dispose of the phagosomal contents. This aspect

of the phagosomal life cycle—referred to here as resolution—is by far the least understood.

1.2.2 Phosphoinositides in phagocytosis

When phagocytic targets are initially encountered, extracellular signals must be conveyed

across the plasma membrane (PM) in order to initiate the complex cellular behaviors that

culminate in phagocytosis. It is becoming increasingly apparent that phosphoinositides play a

prominent role in relaying this information. Indeed, both the ruffling of membranes for probing,

and the detection of ligands by phagocytic receptors are accompanied by local changes in

phosphoinositide composition. Similarly, phosphoinositides coordinate membrane fusion and

fission events that lead to the acquisition of lysosomal properties during the course of

maturation (Vieira et al., 2001). This is accomplished primarily via recruitment of effector

proteins by a combination of stereochemical and electrostatic interactions. Thus,

phosphoinositides are much more than mere building blocks or structural bystanders of cellular

4

membranes. Instead, they fine-tune signal transduction pathways (Di Paolo and De Camilli,

2006), help specify organelle identity (Kutateladze, 2010), and direct membrane traffic

(Simonsen et al., 2001). Given its pivotal role in phagocyte function, phosphoinositide

metabolism is often subverted by pathogens as an invasion or colonizing strategy. By

undermining or hijacking phosphoinositide homeostasis, intracellular bacteria alter membrane

dynamics and gain entry into their host (Ham et al., 2011). Maturation of the bacterium-

containing vacuole into an organelle with lysosomal characteristics is similarly affected by the

subversion of phosphoinositide signaling.

In this chapter, first I will focus on those events that must occur for phagocytosis to achieve its

ultimate goal: the maintenance of organismal homeostasis through the internalization,

inactivation and disposition of external particles. Initially, I address the events that must occur

for particle engagement and productive signaling allowing for internalization. Then, I detail the

maturation steps that confer upon the phagosome its potent degradative capacity. Lastly, I

define the resolution step the least understood of the phagocytic stages. Where appropriate, I

have extrapolated knowledge acquired studying other endocytic processes to fill the gaps in the

understanding of phagocytosis. Secondly, I collate the available information regarding the

involvement of phosphoinositides in phagocytosis and macropinocytosis. This section also

features selected examples of the molecular mechanisms by which bacterial pathogens

commandeer phosphoinositide homeostasis, thereby compromising phagocytic defenses.

1.3 Stage 1: Phagosome formation

1.3.1 Engaging the target

For phagocytosis to occur, a phagocyte must be in direct physical contact with the target. In

some instances this can occur by passive means, as is the case for senescent red blood cells

that are brought into contact with splenic macrophages by the vascular flow (Kohyama et al.,

5

2009). In the case of some pathogens, thermal forces along with flagellar and ciliary movement

keep phagocytic targets in a constant state of motion, resulting in occasional and fleeting

contact with the surface of host cells (Bray, 2001). More frequently, though, phagocytes must

actively survey their environment for phagocytic targets.

Neutrophils, for example, recognize complement-opsonized particles from a distance, and

migrate up a chemotactic concentration gradient generated by complement by-products to

engage and internalize the targets (Lee et al., 2011). Similarly, both neutrophils and

macrophages migrate along N-formyl-methionyl-leucyl-phenylalanine (fMLP) gradients to

pursue and eventually ingest bacteria (Heit et al., 2008; Devosse et al., 2009). During

efferocytosis—the phagocytosis of apoptotic cells—‘find-me’ signals such as sphingosine 1-

phosphate and fractalkine are released by dying cells that attract macrophages to effect

clearance (Gude et al., 2008; Truman et al., 2008). Given that there are approximately 30

trillion cells in the human body (Bianconi et al., 2013; Sender et al., 2016) and less than one

percent of those are phagocytes (Freitas, 2000), their ability to migrate and scavenge particles

over a large area is critical.

In addition to migration, phagocytes—particularly macrophages and dendritic cells—display a

rather unique behavior: they continuously extend membrane protrusions (Fig. 1.1a). This

constitutive probing or ‘ruffling’ is distinct from the more dramatic bursts of membrane protrusion

that occur upon exposure to growth factors or other stimuli, like lipopolysaccharides (LPS)

(Koivusalo et al., 2010; Yoshida et al., 2015; Canton et al., 2016). The continuous and dynamic

extension of protrusions enhances the capacity of phagocytes to detect and capture stationary

or randomly moving particles (Patel and Harrison, 2008; Flannagan et al., 2010). The force that

drives the extension of cell surface protrusions is largely generated by the polymerization of

branched actin networks directly under the PM (Fig. 1.1a) (Campellone and Welch, 2010).

These networks are generated by the actin-related protein 2 and 3 (Arp2/3) complex, a seven-

6

subunit protein complex consisting of ARPC1-5 and Arp2 and 3, now recognized to have an

added layer of complexity resulting from the existence of multiple isoforms of some subunits

(Mullins et al., 1998; Amann and Pollard, 2001; Abella et al., 2016). Arp2/3 complex

involvement in constitutive membrane ruffling in phagocytes is well established (Vargas et al.,

2016). However, the Arp2/3 complex is intrinsically inactive and must be activated by a

nucleation-promoting factor (NPF) (Welch et al., 1998; Goley et al., 2004; Kreishman-Deitrick et

al., 2005; Rodal et al., 2005; Zencheck et al., 2009). The NPFs responsible for constitutive

membrane ruffling have not been definitively established. One such NPF, the Wiskott-Aldrich

syndrome protein (WASP), is often cited as being required for constitutive membrane ruffling in

dendritic cells (Vargas et al., 2016); however, WASP-deficient dendritic cells show no defect in

ruffling (Pulecio et al., 2008). Other NPFs, such as the WASP-family verprolin homolog (WAVE)

isoforms, have been implicated in stimulus-induced membrane ruffling in both non-

hematopoietic and hematopoietic cells (Innocenti et al., 2005; Abou-Kheir et al., 2008; Isogai et

al., 2015), but a definitive role in the constitutive ruffling of phagocytes has not yet been

established. Recently, a G protein-coupled receptor (GPCR), the calcium-sensing receptor

(CaSR), was shown to be required for the constitutive ruffling of macrophages and dendritic

cells (Canton et al., 2016). CaSR signals the robust generation of phosphatidylinositol 3,4,5-

trisphosphate [PtdIns(3,4,5)P3] and phosphatidic acid (PtdOH) at the PM, which can in turn

recruit the WAVE complex (Oikawa et al., 2004; Suetsugu et al., 2006) and Rac-specific

guanine nucleotide-exchange factors (GEFs) such as DOCK1 (Bohdanowicz et al., 2013;

Sanematsu et al., 2013). Rac can in turn activate both

7

Figure 1.1. Phagosome formation. The events leading to phagosome formation can be summarized in four discrete steps: dynamic probing, particle binding that induces receptor clustering, phagocytic cup formation, and phagosome sealing. a. Macrophages continuously extend membrane protrusions to engage phagocytic targets. Protrusions are driven by Arp2/3-dependent branched actin networks underneath the PM. The Arp2/3 complex is activated by both PtdIns(4,5)P2 and Rho-family GTPases, specifically Cdc42 or Rac. The GTPases are activated locally by GEFs, which are themselves recruited by PtdOH and PtdIns(3,4,5)P3 at the PM. Signals from GPCRs, such as CaSR, induce signaling cascades for protrusion formation. b. In their un-clustered state, phosphorylation and therefore activation

8

of phagocytic receptors is minimized by the activity of phosphatases and by phosphorylation of Src-family kinases by Csk at an inhibitory site. Clustering of phagocytic receptors results in the physical exclusion of large transmembrane phosphatases from sites of receptor engagement. Phosphorylation of phagocytic receptor ITAMs culminates in the activation of Rho-family GTPases. c. The activation of Rho-family GTPases activates NPFs, allowing for the actin-driven extension of the plasma membrane around the phagocytic target. BAR domain-containing proteins may also be involved in NPF activation. The PLCγ-dependent generation of a cytosolic Ca2+ transient results in the activation of myosin, which may facilitate phagosome cup formation and sealing. d. By analogy with other forms of endocytosis, BAR domain-containing proteins may facilitate phagosomal sealing by allowing for directed actin polymerization and for the membrane deformation required for the close apposition of membranes at sites of sealing. In addition, BAR domain-containing proteins can recruit proteins involved in scission, such as dynamin. Components delimited by dashed lines and with higher transparency, e.g. Hs1 in panel d., designate extrapolations from other systems that have not been directly validated in phagocytic systems.

WASP and the WAVE complex to generate branched actin networks (Fig. 1.1a) (Campellone

and Welch, 2010).

The active probing described above brings potential targets in contact with the phagocyte

surface, where an impressive array of receptors is expressed. These include receptors like

TIM4 and MerTK that recognize homeostatic debris typified by apoptotic cells (Flannagan et al.,

2014; Lew et al., 2014), and also receptors such as MARCO (macrophage receptor with

collagenous structure) that bind bacteria (Novakowski et al., 2016). Importantly, not all

phagocytes are ‘created equal’: different phagocytes express distinct but overlapping sets of

phagocytic receptors, bestowing upon them the capacity to recognize specific targets. A

pertinent example is the differential expression of receptors on differentially polarized

phagocytes. The ability of phagocytes to adapt to environmental cues by assuming distinct

functional phenotypes has gained increasing attention in recent years. This capacity, referred to

as polarization, results in drastic differences in the expression of phagocytic receptors. Anti-

inflammatory macrophages (referred to as M2), for example, express efferocytic receptors such

as MerTK and CD36 at much higher levels than inflammatory macrophages (referred to as M1)

(Zizzo et al., 2012; Huang et al., 2014). M1 macrophages, on the other hand, express higher

levels of activating Fcγ receptors (Beyer et al., 2012) and lower levels of the inhibitory FcγRIIb

(Pricop et al., 2001). As a result, M1 macrophages internalize immunoglobulin G (IgG)-

9

opsonized targets much more effectively than M2 cells (J. Canton, unpublished observations).

Differential expression of receptors ensures that targets are preferentially engulfed by

phagocytes adept at processing them. In line with this, M2 phagosomes undergo rapid

acidification and maturation, ideal for digesting and recycling apoptotic cells, whereas M1 cells

produce abundant reactive oxygen species (ROS) for the rapid killing of IgG-opsonized

pathogens (Canton, 2014; Canton et al., 2014). Indeed, the aberrant sorting of apoptotic cells

to M1 macrophages was shown to have deleterious implications, including the development of

autoimmunity (Uderhardt et al., 2012).

Not all particles that are contacted by the phagocytes are destined for ingestion. Phagocytic

cells undertake sophisticated information processing to determine whether a particle is to be

engulfed or not. To this end, in addition to the bona fide phagocytic receptors that transduce

signals resulting in particle uptake, phagocytes express various ancillary receptors that convey

information on the nature of the target engaged. In some cases, this additional information can

antagonize phagocytosis or affect the downstream processing of the phagocytic target

(Mantegazza et al., 2014; Nair-Gupta et al., 2014).

1.3.2 Transducing the signal

Once contact is made with the phagocytic target, the phagocytic and ancillary receptors initiate

a signaling cascade. Whether or not the particle is to be ingested depends on the type and

density of ligands engaged. In some instances, in addition to phagocytic determinants the

particles harbor ‘don’t-eat-me’ signals that exert a potent inhibitory effect, precluding engulfment

(Arandjelovic and Ravichandran, 2015). The biophysical and molecular requirements for the

productive signaling of phagocytosis are considered in discrete steps in this section.

10

i) Lateral diffusion of receptors. A common feature of many phagocytic receptors is the

requirement for lateral clustering for signal initiation. Several phagocytic receptors, including

dectin-1 and the Fc receptors, associate with and can be phosphorylated by Src-family tyrosine

kinases (SFKs). They rely on clustering to elevate the local concentration of substrate and

kinase (Flannagan et al., 2012; Underhill and Goodridge, 2012) and also for the physical

exclusion of phosphatases (Fig. 1.1b) (Goodridge et al., 2011; Yamauchi et al., 2012; Freeman

et al., 2016). The ability of phagocytic receptors to move laterally in the plane of the membrane

and hence their propensity to cluster underneath multivalent ligands is directly influenced by the

cytoskeleton (Jaqaman et al., 2011; Jaumouillé et al., 2014). Single-particle tracking revealed

that transmembrane receptors, including the phagocytic receptors FcγR and SR-B2, do not

undergo simple Brownian diffusion in the plane of the PM (Kusumi et al., 2005; Jaqaman et al.,

2011; Jaumouillé and Grinstein, 2011; Trimble and Grinstein, 2015; Fujiwara et al., 2016).

Rather, they display restricted motion within defined regions of the PM (Kusumi et al., 2005).

Regions of confinement are delineated by an actin meshwork that generates ‘fences’ (Kusumi et

al., 2014; Fujiwara et al., 2016); the ‘pickets’ of such fences—transmembrane proteins anchored

to the actin network—exert steric and viscosity effects that hinder the movement of

transmembrane receptors (Fujiwara et al., 2016). For some phagocytic receptors, specifically

integrins, indirect linkage to the actin meshwork can also limit mobility (Kanchanawong et al.,

2010). These regions of confinement can restrict the initiation of phagocytosis in several ways:

a) by curtailing the lateral diffusion and hence the clustering of phagocytic receptors; b) by

decreasing the overall avidity of target binding by restricting the access of receptors to their

ligands and c) by preventing the interaction of phagocytic receptors with activating/synergistic

co-receptors that may exist in discrete membrane subdomains. Therefore, cytoskeletal

rearrangement conducive to receptor clustering is a pre-requisite for efficient signal initiation

during phagocytosis (Jaumouillé et al., 2014). In line with this, macrophages display a degree

of tonic signaling by spleen tyrosine kinase (Syk) and SFKs, which maintains a more open and

11

dynamic actin meshwork that facilitates lateral clustering upon receptor engagement

(Jaumouillé et al., 2014). Moreover, large scale Syk-dependent actin rearrangement near the

tips of advancing pseudopods during phagocytosis fosters increased lateral mobility and

clustering of receptors (Jaumouillé et al., 2014). It is very likely that non-phagocytic accessory

receptors also rearrange the actin meshwork to modulate receptor mobility and therefore

phagocytic efficiency. Toll-like receptor 4 (TLR4) signaling in B cells, for example, activates the

actin-severing protein cofilin, thereby lowering the threshold for activation of the B cell receptor

(Treanor et al., 2010; Mattila et al., 2013; Freeman et al., 2015). In macrophages, TLR4

signaling is known to enhance phagocytosis of several targets (Blander and Medzhitov, 2004;

Underhill and Gantner, 2004; Anand et al., 2007; Kong and Ge, 2008; Chen et al., 2012); this

effect is likely due, at least in part, to a similar actin rearrangement-dependent mechanism. G

protein-coupled receptors such as CX3CR1 can also prime phagocytosis, an effect associated

with and attributable to dynamic rearrangements of the cortical cytoskeleton (Borgman et al.,

2014; Wong et al., 2016).

ii) Initiation of receptor signaling. Several bona fide phagocytic receptors, such as the FcγRs,

possess an immunoreceptor tyrosine-based activation motif (ITAM) in their cytosolic domain.

ITAMs are found on the accessory Fc receptor γ-chain in FcγRI and FcγRIIIa and on the α chain

in FcγRIIa/c (Guilliams et al., 2014). They consist of the conserved sequence YxxL/I repeated

twice and typically spaced by about six to twelve amino acid residues. Variations of the ITAM,

such as the hemi-ITAM of dectin-1 also exist (Underhill and Pearlman, 2015). ITAM and hemi-

ITAM motifs are phosphorylated by dedicated tyrosine kinases, notably the SFKs (Fig. 1.1b)

(Fitzer-Attas et al., 2000) and also Syk. Phosphorylation is markedly stimulated upon receptor

clustering, but can in principle happen also spontaneously, as a result of fortuitous collision

between receptors. To prevent untimely activation, negative regulators exist (Fig. 1.1b). These

include C-terminal Src kinase (Csk) that keeps SFKs inactive by phosphorylating an inhibitory

tyrosine residue near their C-terminus (Martin, 2001; Dodd et al., 2014; Freedman et al., 2015),

12

and also tyrosine phosphatases. Productive signaling only occurs when the basal inhibition

exerted by these phosphatases is overcome. Phagocytic receptor clustering achieves this by at

least two mechanisms: the local accumulation of activating kinases and the physical exclusion

of inhibitory phosphatases.

Curiously, conditions that result in the clustering of activating FcγRs can also co-aggregate

inhibitory FcγRs, such as FcγRIIb, which have an immunoreceptor tyrosine-based inhibition

motif (ITIM). ITIMs are also phosphorylated by SFKs (Malbec et al., 1998) and can recruit the

SH2-containing tyrosine phosphatases SHP-1 and SHP-2 (Famiglietti et al., 1999). SHP-1 and

SHP-2 can in turn dephosphorylate nearby ITAMs (Getahun and Cambier, 2015). It is of

interest that ITIM-bearing FcγRs can be partially excluded from sites of phagocytic target

contact on the basis of differential affinity for the IgG isotype opsonizing the phagocytic target

(Syam et al., 2010).

iii) Productive receptor signaling. The local concentration of active SFKs at sites of receptor

clustering, together with the exclusion of countervailing phosphatases, results in the

phosphorylation of the activating tyrosine residues in the receptor ITAM (or hemi-ITAM). The

phosphorylated tyrosines in turn serve as a docking site for the tandem SH2 domains of Syk

(Fig. 1.1) (Johnson et al., 1995). Syk activity is required for FcγR-mediated phagocytosis,

although its role in dectin-1 mediated phagocytosis is less clear (Crowley et al., 1997; Kiefer et

al., 1998; Herre et al., 2004; Rogers et al., 2005; Underhill et al., 2005; Jaumouillé et al., 2014;

Deng et al., 2015). Syk amplifies signaling by further phosphorylating nearby ITAMs (Mócsai et

al., 2010), but also propagates the signal through the recruitment and phosphorylation of

adaptor proteins. Several adaptor proteins including linker of activated T cells (LAT), growth

factor receptor-bound protein 2 (Grb2), Grb2-associated-binding protein 2 (Gab2), and the Src

homology 3 (SH3) domain-binding protein 2 (SH3BP2) are recruited to sites of phagocytic

receptor signaling in a Syk-dependent manner (Fig. 1.1b) (Tridandapani et al., 2000; Yu et al.,

13

2006; Prod’Homme et al., 2015). It is these adaptors that then recruit cytosolic effectors to carry

out the extensive lipid and cytoskeletal remodeling that accompanies the phagocytic process.

Local changes to the lipid composition of the inner leaflet of the PM at sites of phagocytic

receptor clustering serve to spatially and temporally coordinate phagocytic signaling. Entire

sections of this chapter are dedicated to comprehensively describe these paramount events.

1.3.3 Cytoskeletal rearrangements

The signaling described above culminates in the extension of actin-driven projections of the PM

that advance around phagocytic targets, eventually sealing at their distal tips to form a

phagosome. Although the mechanisms that drive actin polymerization around phagocytic

targets vary depending on the nature of the target/opsonin, the general requirements remain the

same. Specifically, there is the activation and recruitment of an NPF, followed by the Arp2/3-

dependent nucleation of branched actin networks (Fig. 1.1c). Of the NPFs capable of activating

Arp2/3, WASP and N-WASP have the most clearly defined role in several modes of

phagocytosis, including both integrin- and FcγR-mediated phagocytosis (Lorenzi et al., 2000;

May et al., 2000; Park and Cox, 2009). A number of mechanisms account for the recruitment

and activation of WASP/N-WASP to sites of phagosome formation. An increase in

PtdIns(4,5)P2 levels at the site of engagement is likely to play a role, this will be extensively

described in a subsequent section of this chapter. Conversion of Cdc42 to its GTP-bound state

at sites of phagosome formation occurs, at least in part, through the recruitment of the GEF

intersectin-1 by a complex consisting of WASP/N-WASP and the adaptor proteins Nck and Grb2

(Dart et al., 2012; Humphries et al., 2014). This signaling node is further buttressed by the

interactions of the SH2 and SH3 domains of Nck with phosphorylated ITAMs and poly-proline

motifs in WASP/N-WASP, respectively (Dart et al., 2012; Humphries et al., 2014). The

coordination of Cdc42 and WASP/N-WASP during phagosome formation may also be regulated

14

by members of the Bin-Amphiphysin-Rvs (BAR) domain-containing family of proteins that bind

to sites of membrane curvature (Fig. 1.1c). Accordingly, macrophages deficient in the Fer and

Cip4 homology-BAR (F-BAR) protein FBP17, which has a Cdc42-binding site and an SH3

domain capable of binding WASP/N-WASP, have defects in phagocytic cup formation (Tsuboi

et al., 2009). Similarly, deficiency of the inverse-BAR (I-BAR) protein IBARa in Dictyostelium

spp., results in defective phagocytosis (Linkner et al., 2014).

It is somewhat surprising that activation of Arp2/3 via the WAVE complex does not seem to be

required for phagocytosis (Kheir et al., 2005). The WAVE complex is known to be recruited to

the PM at sites of PtdIns(3,4,5)P3 accumulation through a basic motif and also interacts with

Nck via Nck-associated protein 1 (Nap1) (Campellone and Welch, 2010). Additionally, a

deficiency in the small GTPase Rac, which is an upstream activator of the WAVE complex, is

associated with defective actin polymerization at sites of phagosome formation (Cox et al.,

1997; Koh et al., 2005; Hall et al., 2006; Utomo et al., 2006). Based on these facts, one would

anticipate WAVE to contribute significantly to actin assembly during phagocytosis. However,

Rac can also activate N-WASP by binding to its Cdc42- and Rac-interacting binding (CRIB)

motif (Tomasevic et al., 2007). Moreover, Rac is also involved in the activation of PIP5K to

produce PtdIns(4,5)P2, which enhances the recruitment of WASP/N-WASP to the PM (Weernink

et al., 2004). In conclusion, the elaboration of branched actin networks during phagocytosis is

driven by Arp2/3-dependent nucleation through the local recruitment of NPFs. These events

are spatially and temporally regulated by signaling adaptors downstream of ITAM

phosphorylation, and by the generation of crucial lipid intermediates at sites of receptor

engagement.

15

1.3.4 Sealing the phagosomal membrane

Perhaps the most enigmatic event during phagosome formation is the coalescence of the

membrane protrusions at their distal margins to form a sealed phagosome. The gap in

understanding phagosomal sealing has been tentatively filled by insights gained studying other

endocytic processes. For fusion to occur, the protruding membrane margins must initially be

brought into contact. As such, the force generated by the advancing actin protrusions must

have directionality. In other modes of endocytosis, directionality is achieved by the binding of

BAR domain-containing proteins to sites of negative curvature.

A prototypical example is the constriction of the neck of a forming clathrin-coated pit (CCP).

During CCP formation, BAR domain-containing proteins of increasing curvature are recruited to

sites of neck constriction (Daumke et al., 2014). Due to their more relaxed curvature, the F-

BAR proteins, such as FBP17, are recruited to sites of PtdIns(4,5)P2 enrichment early in pit

formation (Itoh et al., 2005). FBP17 facilitates Arp2/3-dependent nucleation directed towards

the site of FBP17 binding (Tsujita et al., 2006; Takano et al., 2008). At later stages of CCP

formation, N-terminal amphipathic helix-containing BAR (N-BAR) proteins, which have greater

curvature, are recruited to sites of neck constriction (Daumke et al., 2014). Like the F-BAR

proteins, N-BAR proteins such as endophilin and amphiphysin also facilitate Arp2/3-dependent

nucleation and additionally recruit dynamin, a GTPase involved in scission (Kaksonen et al.,

2006). This complex is further stabilized through interactions of the proline-rich domain of

dynamin with the SH3 domain of cortactin—or its homolog, hematopoietic lineage cell-specific

protein 1 (Hs1)—which in turn stabilizes Arp2/3 branch points and recruits WASP/N-WASP,

thereby facilitating Arp2/3-dependent actin polymerization (Buccione et al., 2004). Altogether,

the sequential recruitment of BAR domain-containing proteins coordinates actin-dependent

membrane invagination, neck constriction and ultimately recruitment of the scission machinery

during clathrin-mediated endocytosis (Buccione et al., 2004; Kaksonen et al., 2006; Mooren et

16

al., 2012; Daumke et al., 2014). Of note, similar events do occur during phagocytosis. FBP17

and the N-BAR protein bridging integrator-2 (Bin2) are recruited to phagocytic cups, where they

regulate phagocytic efficiency (Tsuboi et al., 2009; Sánchez-Barrena et al., 2012). The N-BAR

protein amphiphysin also mediates actin polymerization during phagocytosis (Gold et al., 2000;

Yamada et al., 2007). Finally, dynamin-2 is required for the extension of membrane protrusions

around phagocytic targets, as well as for the final scission step (Gold et al., 1999; Tse et al.,

2003; Otsuka et al., 2009; Marie-Anaïs et al., 2016). It is therefore tempting to suggest that

BAR domain-containing proteins serve similar functions in CCP and phagosome formation,

namely coordinating actin polymerization and ultimately scission (Fig. 1.1d). When drawing

analogies between CCP and phagosome formation, however, it is important to consider that the

margins of the developing phagosomal cup—which need to be brought together—are separated

by a comparatively enormous distance. This has given rise to the notion of a ‘purse string’

effect, whereby a contractile force draws the phagosomal borders closer (Fig. 1.1d).

Constriction of deformable phagocytic targets has been reported, giving credence to the

existence of a contractile force. In this regard, the activity of myosin isoforms IC, II and X, has

been invoked to account for neck constriction at sites of phagosomal sealing (Swanson et al.,

1999; Cox et al., 2002; Araki et al., 2003).

1.3.5 Terminating the signal

To complete the phagocytic process, the signals that dictate the formation of actin-driven

protrusions must be terminated. SH2 domain-containing tyrosine phosphatases, such as SHP-1

and SHP-2, are recruited to sites of phagosome formation (Ganesan et al., 2003), where they

dephosphorylate and inactivate ITAMs as well as phosphorylated SFKs, Syk and PI3K

(Flannagan et al., 2012). The PtdIns(3,4,5)P3 phosphatase SHIP is similarly attracted to the

cup. However, it is unclear whether these events are intended to terminate phagocytosis, or to

moderate its intensity. Indeed, live-cell imaging has revealed that termination is a much more

17

complex, exquisitely choreographed event: even as actin polymerizes at the tip of advancing

pseudopods, it is simultaneously depolymerizing at the base of the phagocytic cup. This

delicate spatial and temporal coordination is particularly important for the internalization of large

particles (Araki et al., 1996; Cox et al., 1999a). Depolymerization at the base of forming

phagosomes may serve to deliver rate-limiting components of the actin polymerization

machinery to the leading edge of pseudopods. It may also remove a physical barrier to the

exocytosis of endomembranes that need to fuse with the base of the cup, and/or may allow for

membrane deformation required for sealing and scission.

Just as in actin polymerization, lipid-remodeling events also drive depolymerization. As

mentioned earlier, these events will be described in detail in further sections dedicated to lipids

during phagocytosis.

1.4 Stage 1: Phagosome maturation

Phagosome formation, albeit remarkably specialized and complex, is only the first step in

achieving the ultimate goal of phagocytosis: the inactivation and degradation of the engulfed

material. To this end, the membrane and luminal contents of the newly formed phagosome

must undergo a drastic transformation. This gradual conversion, known as phagosome

maturation, generates a hostile, degradative environment that causes the destruction of the

phagocytic prey. In fact, several pathogens have evolved diverse strategies to evade

phagosome maturation, emphasizing its importance in immunity. The substantial number of

different mechanisms used by pathogens to thwart maturation, avoid killing or escape the

phagosome altogether have been reviewed (Flannagan et al., 2012). In the process, specific

components of the phagosome must be preserved and repurposed for diverse cellular functions.

The following section describes the events governing phagosome maturation, their ostensible

purpose and underlying molecular mechanism. By analogy with endosomal compartments,

18

phagosome maturation is divided into sub-stages: the early phagosome, the late phagosome

and the phagolysosome. These are described separately below. Maturation has been

analyzed in most detail in macrophages and the results of these studies constitute much of the

information summarized in this section. However, it is clear that phagosome maturation varies

drastically among phagocytic cell types. For this reason, a specific sub-section is devoted to

illustrate the differences between macrophages and neutrophils during the degradative stage.

1.4.1 The early phagosome

Technically, maturation begins upon phagosome scission from the PM. However,

endomembranes fuse with the forming phagosome even before its closure (Bajno et al., 2000;

Bohdanowicz et al., 2012a). Once internalized, the newly formed phagosome continues to

undergo fusion events, initially with early endosomes. Unlike the bulk PM that it derives from,

the phagosome must be ‘primed’ for fusion, but the details of such priming are poorly

understood.

A pivotal event in phagosome maturation is the acquisition of active Rab5, a GTPase that is

necessary to promote early fusion events (Roberts et al., 2000; Vieira et al., 2003). A number of

GEFs have been implicated in Rab5 activation. One such GEF is GAPex-5, which has been

linked to maturation following the phagocytosis of apoptotic cells (Kitano et al., 2008). Evidence

from endosomal systems suggests that Rab22a, which is also detected in phagosomes

(Roberts et al., 2006), may also be important for Rab5 activation. Rab22a recruits the Rab5

GEF, Rabex-5, promoting Rab5 activation (Fig. 1.2) (Zhu et al., 2009). Rabaptin-5, a Rab5

effector, stimulates Rabex-5 activity resulting in a positive-feedback loop for Rab5 activation

(Horiuchi et al., 1997; Lippé et al., 2001; Zhang et al., 2014).

19

Figure 1.2. Early maturation: from the new phagosome to the early phagosome. Maturation begins as the newly formed phagosome becomes an early phagosome. Center: The newly formed phagosome (left half), still enriched in PtdIns(3,4,5)P3, rapidly loses its plasma membranes markers. The new phagosome undergoes fusion with early endosomal compartments, giving rise to the early phagosome (right half). The early phagosome is enriched in PtdIns3P and its lumen is moderately acidic. Top left: For activation of the small GTPase Rab5, GTP-bound Rab22a can recruit Rabex-5, a Rab5 GEF. Upon activation, Rab5 recruits its effector Rabaptin 5, which in turn stimulates Rabex-5 activity for a positive feedback loop for Rab5 activation. Middle left: The Rab5 effector Vps34 generates PtdIns3P from PtdIns. Bottom left: Together, Rab5 and PtdIns3P recruit EEA1 that stimulates fusion of the phagosome with early endosomes by docking membranes and interacting with SNAREs. Active Rab5 recruits the membrane-tethering CORVET complex. The complex also interacts with SNAREs proteins. Bottom right: Rab11 and Rab4 are involved in recycling traffic to the plasma membrane, the Rab11 effector RCP has been proposed as a recycling regulator from the phagocytic compartment. Middle right: Early phagosomes recruit SNXs through their PX domains. SNXs bear BAR domains capable of inducing, sensing and/or stabilizing membrane curvature. The retromer, comprised of a SNX dimer (SNX1 or SNX2 and SNX5 or SNX6; recruited to the early phagosome) and a cargo-recognition trimer (Vps26,

20

Vps29 and Vps35; recruited to the late phagosome) mediates retrograde traffic to the TGN through tubulovesicular structures. Top right: Ubiquitinated cargo, such as FcγR, is recognized by ESCRT-0. Then, ESCRT-I, -II and –III are sequentially recruited to the membrane, mediating its invagination. This results in the formation of intraluminal vesicles that are targeted for degradation.

Active Rab5 promotes membrane fusion events in several ways. One of the crucial Rab5

effectors is the PI3K Vps34 (Christoforidis et al., 1999b; Vieira et al., 2001; Kinchen et al.,

2008). The product of Vps34 activity is PtdIns3P (Stack and Emr, 1994; Stephens et al., 1994),

the characteristic phosphoinositide of the early phagosome (Fig. 1.2) (Vieira et al., 2001).

PtdIns3P is central to numerous events during early phagosome progression that will be

discussed later in this chapter. The recruitment of the early endosomal antigen 1 (EEA1) is an

example. Rab5 and PtdIns3P synergize to recruit the EEA1 to the phagosomal membrane (Fig.

1.2) (Simonsen et al., 1998; Lawe et al., 2002). EEA1 facilitates fusion by docking target

membranes (Fig. 1.2) (Christoforidis et al., 1999a) and interacting with the fusion-mediating

soluble NSF attachment protein receptors (SNAREs) Syntaxin 6 (Simonsen et al., 1999) and

Syntaxin 13 (McBride et al., 1999). Blocking EEA1 with microinjected antibodies impairs

phagosome maturation (Fratti et al., 2001a).

In yeast endosomes, the class C core vacuole/endosome tether (CORVET) complex facilitates

fusion by tethering membranes from different compartments. The complex, which consists of

four core subunits (Vps11, Vps16, Vps18 and Vps33) and two specific subunits (Vps3 and

Vps8), is another Rab5 effector (Fig. 1.2) (Peplowska et al., 2007). Of note, the core subunits

are shared between CORVET and the homotypic fusion and vacuole-sorting (HOPS) complex,

involved in late endosomal fusion. GTP-bound Rab5 specifically binds Vps3 and Vps8 (Epp

and Ungermann, 2013). Moreover, Vps33 can bind soluble SNAREs to promote fusion (Fig.

1.2) (Lobingier and Merz, 2012). While there are as yet no reports of the recruitment or activity

of the CORVET complex in phagosomes, its known interactions with Rab5 make it a likely

candidate for tethering of early endosomes to phagosomes. Additionally, in yeast, CORVET

21

was implicated in the transition from Rab5 to Rab7, a critical event during the maturation of the

phagosome discussed later in this chapter.

Although the ultimate purpose of the phagosome is, ostensibly, to degrade its contents, specific

phagosomal constituents have diverse fates. While the cargo is targeted for degradation,

several components are recycled to the PM or through retrograde transport to the Trans Golgi

Network (TGN). Because a substantial portion of the PM can be internalized during

phagocytosis, mechanisms must exist in order to replenish it. Soon after phagosome formation,

proteins such as the transferrin receptor are rapidly shuttled back to the PM (Fig. 1.2) (Buys and

Kaplan, 1987). To carry out this segregation, the phagosome must acquire specialized

machinery. In this context, it is interesting that Vamp3-positive recycling endosomes are

delivered focally in an Arf6-dependent manner to the forming phagosome before its closure

(Bajno et al., 2000; Niedergang et al., 2003). Recycling events are primarily mediated by Rab

GTPases, specifically Rab11 (slow recycling) and Rab4 (fast recycling) (Fig. 1.2). Both proteins

have been detected in proteomic analyses of isolated phagosomes (Garin et al., 2001).

Moreover, when overexpressed, Rab11 localizes to the early phagosome, while expression of a

GTP-binding-deficient Rab11 impairs FcγR-mediated phagocytosis (Cox et al., 2000). The

Rab11 effector Rab Coupling Protein (RCP), which is found in the early phagosome, was

proposed to regulate recycling (Fig. 1.2) (Damiani et al., 2004).

In addition to outward vesiculation, the limiting membrane of the phagosome also undergoes

inward budding. This results in the formation of intraluminal vesicles (ILVs) (Lee et al., 2005)

whose membrane and cargo are targeted for degradation. The endosomal sorting complex

required for transport (ESCRT) drives inward vesiculation. ESCRT consists of four complexes

(ESCRT-0, ESCRT-I, ESCRT-II and ESCRT-III) that act sequentially (Fig. 1.2). Ubiquitinated

cargo, such as FcγR (Booth et al., 2002), is recognized by components of the complex (Wollert

and Hurley, 2010). Additionally, ESCRT-0 is recruited to the phagosome, at least in part, via its

22

Hrs subunit, that binds to PtdIns3P through its FYVE domain (Vieira et al., 2004). After

invagination occurs, ESCRT-III mediates scission from the limiting phagosomal membrane (Fig.

1.2) (Olmos and Carlton, 2016). The resulting ILVs are then degraded as the phagosome

transitions to the late phagosome and phagolysosome stages. Notably, other mechanisms

involving ceramide (Trajkovic et al., 2008) and lysobisphosphatidic acid (Matsuo et al., 2004)

have been reported to contribute to multivesicular body formation in the endosomal pathway

and may play a similar role in phagosome maturation.

The sorting capacity of the phagosome relies in part on its ability to deliver elements to the

TGN. For this, the phagosomal membrane undergoes another type of pronounced deformation.

Retrograde traffic to the TGN occurs via tubulovesicular structures similar to those reported in

the endocytic pathway (Hierro et al., 2007). Tubulation involves recruitment of sorting nexins

(SNXs), a family of proteins that bear PX and BAR domains. Some SNXs are recruited to

phagosomes in a PtdIns3P-dependent fashion through their PX domain (Fig. 1.2) (Bonifacino

and Hurley, 2008). Additionally, BAR domains, which can induce, sense and/or maintain

curvature, likely contribute to the generation of vesicles and/or tubules emanating from the

phagosome (Fig. 1.2). Intriguingly, evidence suggests that retrograde transport is completed in

the late maturation stage, as described below.

1.4.2 Early to late phagosome transition

The early phagosome then fuses with late endosomes, thereby becoming a late phagosome.

The late phagosome has a more acidic luminal pH due to the acquisition of additional copies of

the proton-pumping V-ATPase. The late phagosome is also enriched in ILVs. An essential step

during this transition is the ‘switch’ from a Rab5-positive compartment to a Rab7-positive one.

As described before, Rab proteins are activated by GEFs. Though once a subject of debate, it

is now widely accepted that, at least in endosomes, the GEF for Rab7 is Mon1-Ccz1

23

Figure 1.3. Early to late phagosome transition and phagolysosome biogenesis. Center: The early phagosome undergoes fusion with late endosomes giving rise to the late phagosome, a more acidic compartment. Late phagosomes fuse with lysosomes, forming phagolysosome, the ultimate degradative organelles. Top left: The switch from active Rab5 to active Rab7 is likely mediated by the Mon1-Ccz1 complex. Mon1 interacts with Rabex-5 and mediates the displacement of the Rab5 GEF from the membrane. Mon1 then recruits its binding partner, Ccz1 that is thought to facilitate the displacement of the Rab7 GDI. The Mon1-Ccz1 complex functions as the Rab7 GEF. Middle left: The late phagosome acquires in PI4K2A. PtdIns4P, the product of PI4K2A, is present at the late stage. Bottom left: The Rab5 to Rab7 transition mediates the switch from the CORVET complex to the HOPS complex. Four subunits are shared between the complexes; Vps3 and Vps8 (Rab5-interacting subunits) from CORVET are replaced by Vps39 and Vps41 (Rab7-interacting subunits) in HOPS. Bottom center: HOPS mediates tethering between Rab7-positive compartments. Additionally, Vps33 promotes late fusion events through its interactions with SNAREs. The late phagosome acquires LAMP-1 and -2 through fusion with late endosomes and lysosomes. The SNAREs Vamp7, Syntaxin7, Syntaxin8 and likely Vamp8 mediate the biogenesis of the phagolysosome. Bottom right: Rab7 bridges the phagosome with microtubules through its effectors RILP and ORP1L. RILP recruits the dynein-dynactin microtubule motor proteins that direct

24

centripetal migration of the phagosome towards the MTOC. Middle right: The cargo recognition trimer (Vps25-Vps29-Vps36) of the retromer complex binds active Rab7. The complex is necessary for retrieval of M6PR from the phagosome to the Golgi. Top right: The phagolysosome is enriched in hydrolases (proteases, nucleases and lipases) and contains potent antimicrobial peptides. Other crucial components of the phagolysosome are: the V-ATPase (that pumps protons from the cytoplasm to the phagosomal lumen), NOX2 (that transports electrons from cytosolic NADPH to molecular oxygen in the lumen of the phagosome generating ROS) and the voltage-gated proton channel HV1.

(Nordmann et al., 2010). The Rab switch entails two strictly coordinated stages. First, Mon1

interacts with Rabex-5. Upon its recruitment to the transitioning compartment Mon1 displaces

Rabex-5, ending the Rab5 activation positive feedback loop (Fig. 1.3) (Poteryaev et al., 2010).

Interestingly, Mon1 binds PtdIns3P in vitro, hence it has been proposed to sense the

phosphoinositide levels in the maturing compartment. Once recruited, Mon1 binds Ccz1. It is

this dimeric complex that acts as the Rab7 GEF (Fig. 1.3) (Nordmann et al., 2010). Moreover,

evidence suggests that Mon1-Ccz1 promotes Rab7 recruitment to the membrane through the

displacement of the Rab7 GDI (Fig. 1.3) (Kinchen and Ravichandran, 2010).

The transition from Rab5 to Rab7 is the driving force for the CORVET complex to switch and

become the HOPS complex. As mentioned earlier, four common subunits compose the core of

both complexes. Molecularly, the switch consists of displacement of Vps3 and Vps8 subunits of

CORVET on the early compartment—which occurs upon loss of Rab5—followed by their

replacement by Vps39 and Vps41 (Fig. 1.3) (Plemel et al., 2011). The HOPS-specific subunits,

Vps39 and Vps41, bind to membrane-associated Rab7 (Fig. 1.3) (Bröcker et al., 2012). Like

CORVET, HOPS serves as a tether between membranes. Moreover, Vps33, still present in

HOPS, promotes fusion by interacting with SNAREs (Fig. 1.3) (Lobingier and Merz, 2012).

While the role of CORVET in phagosomes is only inferential, there is direct evidence implicating

HOPS in phagosomal systems. Silencing of individual HOPS subunits in C. elegans impairs the

maturation of phagosomes containing apoptotic cells (Kinchen et al., 2008).

25

While retrograde transport from the phagosome to the TGN is initiated during early maturation

(Fig. 1.2) its completion occurs at later stages. This requires the recruitment of a specialized

complex. The full retromer is composed of a SNX dimer (initially recruited by PtdIns3P to the

early phagosome) and the cargo selective trimer of Vps26-Vps29-Vps35 (Figs. 1.2 and 1.3)

(Bonifacino and Hurley, 2008). The trimeric complex associates with active Rab7 (Fig. 1.3),

suggesting that retrograde traffic is completed in the late stage. Accordingly, depletion of Rab7

impairs retromer assembly in endosomes and blocks the retrieval of mannose 6-phosphate

receptor (M6PR) to the TGN (Rojas et al., 2008).

During its maturation, the phagosome undergoes centripetal movement towards the

microtubule-organizing center. GTP-bound Rab7 orchestrates this inward movement. Late

compartments associate with microtubules through two Rab7 effectors: the Rab7-interacting

lysosomal protein (RILP), and the long splice variant of the OSBP-related protein 1 (ORP1L)

(Fig. 1.3) (Harrison et al., 2003; Rocha et al., 2009). With the cooperation of ORP1L, RILP

recruits the dynein-dynactin complex (Fig. 1.3) (Jordens et al., 2001; van der Kant et al., 2013a)

that promotes minus-end transport along microtubules (Fig. 1.3) (Johansson et al., 2007).

Centripetal migration of phagosomes is essential for the generation of phagolysosomes

(Harrison et al., 2003).

Curiously, while PtdIns3P defines the early phagosome, a characteristic lipid of the late

phagosome has not been described. Intriguingly, the PtdIns4P kinase 2A (PI4K2A) (Ketel et al.,

2016) has been found in purified maturing phagosomes (Fig. 1.3) (Jeschke et al., 2015). This

lipid kinase, its product and their functional implications will be discussed in detail in a section

ahead.

26

1.4.3 Phagolysosome biogenesis

The following phagosomal stage, which has been at least partially characterized, is the

formation of the phagolysosome. It is during this stage that the phagosome acquires most of its

degradative components and properties. The fundamental event leading to phagolysosome

biogenesis is the fusion of the late phagosome with lysosomes (Fig. 1.3). As mentioned earlier,

the Rab7-dependent centripetal movement of the phagosome is critical for this fusion event

(Harrison et al., 2003). Additionally, the SNAREs VAMP7, VAMP8, syntaxin 7 and syntaxin 8

have been characterized as important components of the lysosomal fusion machinery (Fig. 1.3)

(reviewed in Luzio et al., 2007). The phagolysosome is enriched in lysosome-associated

membrane proteins 1 and 2 (LAMP-1 and -2) (Fig. 1.3). These integral membrane proteins are

crucial for late maturation stages and microbial killing (Binker et al., 2007; Huynh et al., 2007).

The phagolysosome is highly acidic, oxidative and contains an assortment of lytic enzymes that

make it hostile to the ingested prey. Acidification of the phagosome starts immediately after

sealing and develops gradually, reaching pH values as low as 4.5-5.0 in some cell types.

However, it is worth noting that there is a striking variability in the rate and extent of phagosomal

acidification in different phagocytes (Segal et al., 1981; Lukacs et al., 1991; Hackam et al.,

1997; Mantegazza et al., 2008; Canton et al., 2014). Neutrophils, for example, maintain a

slightly alkaline phagosomal pH for extended periods of time (Jankowski et al., 2002), while

phagosomes in M2 macrophages reach very acidic pH values within minutes of sealing (Fig.

1.4) (Canton et al., 2014). The differential phagosomal pH is reflective of the function of the

particular phagocyte. M2 macrophages are typically involved in the clearance and recycling of

homeostatic debris and, consistent with this, the generation of an acidic pH is required for the

prompt degradation of their phagosomal contents. Indeed, several hydrolytic enzymes acquired

by the phagolysosome have acidic pH optima. Of note, acidification itself seems to promote the

progression of the phagosome. In line with this, dissipation of phagosomal pH gradients was

27

Figure 1.4. Differences in phagosomes between phagocytes. Top panel: The M2 phagolysosome (left side) is rich in active V-ATPases and therefore has an acidic luminal pH (4.5-5.0). In the case of neutrophils and M1 macrophages the pH of the lumen remains close to neutral or becomes alkaline (top panel right side). This is due to the paucity of active V-ATPases in combination with the large rate of proton consumption associated with superoxide dismutation. Bottom panel: Detailed schematic of the contributors to phagosomal pH homeostasis. The V-ATPase is a multi-subunit proton pump. The NADPH oxidase NOX2 is a complex of integral membrane proteins (gp91phox and p22phox) and cytosolic subunits (p40phox, p47phox and p67phox) that are recruited in a Rac1/Rac2-dependent manner. The

28

assembled complex transfers electrons from cytosolic NADPH to luminal oxygen, generating O2-.

Neutrophils and M1 macrophages display higher NOX2 activity than M2 macrophages. The negative charge build-up associated with the electrogenic activity of NOX2 is compensated by the voltage-gated proton channel HV1. The channel allows the flow of protons from the cytosol into the phagosome. Luminal protons and O2

- are used by the SOD to produce H2O2, which can in turn be used for HOCl generation by MPO.

reported to impair the biogenesis of phagolysosomes (Gordon et al., 1980). In contrast to M2

cells, phagosomes of human M1 macrophages, similar to neutrophils and dendritic cells,

maintain pH values close to neutrality (Jankowski et al., 2002; Mantegazza et al., 2008; Canton

et al., 2014). This is a consequence of the production of large quantities of reactive oxygen

species (ROS) and is related to the primary function of M1 macrophages and neutrophils –

killing microbes.

1.5 Stage 3: Phagosome resolution

Phagocytic prey is typically complex and composed of an assortment of macromolecules. In

general, the phagolysosome can catabolize its contents deploying a diverse arsenal of

hydrolytic enzymes in a suitable, pro-degradative environment. However, the products of this

breakdown must be processed to allow the phagocyte to return to homeostasis and to resume

immune responsiveness. Considering that they must dispose of many billions of apoptotic cells

per day, it is evident that phagocytes are extremely efficient waste managers. I therefore felt

that explicit discussion of phagosome resolution was warranted, even though very little is known

about this stage compared to the earlier phagocytic stages. As a result, the following section is

highly speculative, in great part extrapolated from studies on lysosomes and inferred from

lysosomal storage diseases.

29

1.5.1 Disposal of nucleic acids

Various phagocytic targets, including microorganisms, apoptotic bodies and neutrophil

extracellular traps, contain large quantities of nucleic acids. In cells undergoing apoptosis,

nucleic acid degradation is initiated by the action of nucleases within the dying cells (Samejima

and Earnshaw, 2005). This degradation process continues following engulfment by a

phagocyte. Phagocytic cells degrade ingested nucleic acids through DNase II (Fig. 1.5a), a

nuclease delivered to phagosomes upon lysosome fusion (Kawane et al., 2001; Krieser et al.,

2002). The gene encoding DNase II is essential, as DNase II-deficient organisms accumulate

short DNA strands (approx. 200 bp long) in abnormal storage structures (Mukae et al., 2002).

Notably, hematopoietic cells do not generate nucleosides de novo (Hsu et al., 2012). Hence,

they rely on membrane transporters for delivery of extracellular and lysosome-derived

nucleosides to the cytosol for synthesis of nucleic acids. While transport from the extracellular

space has been extensively studied, lysosomal transport, and more particularly phagolysosomal

transport, is less understood. Because nucleoside transporters cannot mediate nucleotide

transport, the latter must first be dephosphorylated. It is conceivable that a nucleotidase carries

out this reaction (Fig. 1.5a). Evidence linking nucleotidases to late phagosomal compartments

is scarce. However, a lysosome-localized 5’-nucleotidase does exist in mammalian cells

(Mellors et al., 1975). Nucleosides can exit lysosomes via the Equilibrative Nucleoside

Transporter 3 (ENT-3), a transmembrane transporter that is highly expressed in macrophages.

Loss of ENT-3 causes a lysosomal-storage disorder in macrophages, characterized by

ineffective apoptotic cell clearance and accumulation of nucleosides after phagocytosis (Hsu et

al., 2012), suggesting a crucial role for this transporter in the resolution of nucleic acid

degradation products (Fig. 1.5a). This sequence of hydrolysis, dephosphorylation and transport

is crucial to the enucleation of mammalian erythrocytes, where the extruded nuclei are directly

engulfed by dedicated macrophages (Manwani and Bieker, 2008).

30

31

Figure 1.5. Phagosome resolution. The major components to be digested and/or recycled during phagocytosis include nucleic acids, proteins, lipids and, eventually, the limiting membrane of the phagosome itself. a) Digestion of nucleic acids is carried out by the luminal enzyme DNase II, yielding nucleotides. Nucleotides are converted into nucleosides by nucleotidases and transported out of the phagosome by nucleoside transporters such as ENT-3. Phagosomes also contain a wide variety of proteases such as the cathepsins that break protein down into peptides and amino acids. Amino acid transporters then translocate the amino acids into the cytosol for reuse or export from the cell. Amino acids can also activate mTORC1 in a mechanism involving the V-ATPase. mTORC1 activation leads to the fission of the phagosome into smaller structures. b) The major lipid constituents to be digested in the phagosome include cholesteryl esters, glycosphingolipids, GM1-2, sphingomyelin and glycerophospholipids. Lysosomal acid lipase converts esterified cholesterol into free cholesterol. Free cholesterol is handed off by NPC2 to NPC1 and subsequently translocated out of the phagosome. ABC transporters similar to those that transport cholesterol at the plasma membrane have been identified in lysosomal compartments, but whether they play a role in the export of cholesterol from the phagosome is unclear. Various lipid carrier proteins, such as the saposins and GM2-activator protein, render glycosphingolipids and gangliosides more accessible to glycosidases for conversion into ceramide. Ceramidase converts ceramide into sphingosine that is transported out of the phagosome, potentially by NPC1. Acid sphingomyelinase also yields ceramide from sphingomyelin. Glycerophospholipids are converted into lysophospholipids by lysosomal phospholipase A2; the mechanism by which lysophospholipids are removed from the phagosome is unclear. c) mTORC1 activation has been linked to phagosomal tubulation and fission into smaller structures. This likely involves kinesin-dependent movement along microtubules, as reported for lysosomes. The accumulation of PtdIns4P on the cytosolic leaflet of the phagosomal membrane may support exocyst-dependent fusion of phagosomal membrane with the plasma membrane.

1.5.2 Protein and amino acid resolution

Virtually every physiological phagocytic target has a significant protein content. As mentioned,

the phagolysosome is enriched in hydrolases, among them proteases, and the luminal

environment is optimal for their action. Cathepsins, an extensively characterized family of

enzymes, are divided into serine, aspartate and cysteine proteases. All of these sub-families

are present in the phagosome and are active during specific maturation stages (Kinchen and

Ravichandran, 2008; Flannagan et al., 2012). Proteases degrade proteins into polypeptides

and further into amino acids that can serve as nutrients for the phagocyte (Fig. 1.5b). This

requires translocation of the amino acids across the membrane, a process facilitated by solute

carriers (SLCs).

32

It is noteworthy that members of the SLC family have recently been implicated in sensing

nutrient availability, delivering the information to enzymes that control anabolic processes

(Dibble and Manning, 2013; Abraham, 2015). The target of rapamycin complex 1 (mTORC1) is

a key element in this sequence; it is found on the cytosolic surface of lysosomes, where it

becomes activated by essential amino acids. SLCs, particularly, SLC38A9, contribute to this

process either directly, by sensing arginine and conveying the information to mTORC1 via the

V-ATPase (Zoncu et al., 2011; Rebsamen et al., 2015; Wang et al., 2015b), or indirectly, by

facilitating the exit of luminal amino acids that then combine with cytosolic Sestrin2, a leucine

sensor that binds and activates mTORC1 (Wolfson et al., 2016). Proteomic analyses have

identified in phagosomes a number of SLCs capable of transporting amino acids (Trost et al.,

2009). It is therefore tempting to speculate that anabolic signals are emitted by

phagolysosomes following the digestion of phagocytic targets, particularly of protein-rich

apoptotic cells.

1.5.3 Lipid processing

Phagosomes have to dispose of engulfed lipids, and the lipid content of physiological

phagocytic targets can be extremely high and diverse. Hence, the phagosome is equipped with

a wide variety of lipases. These enzymes, however, don’t always have access to lipids that are

incorporated into membranes of the prey. In these instances lipids must first be extracted from

bilayers and exposed to the lipases by lipid-transfer proteins, like the saposins.

It is important to note that while the lipids of the prey are targeted for degradation, the limiting

membrane of the phagolysosome must remain intact. Integral membrane (glyco)proteins are

believed to protect the luminal leaflet of the phagosomal membrane from lipolytic enzymes. At a

later stage, however, phagosomes that have completed their degradative function must

themselves be resorbed. To this end, ILVs are formed and degraded in the lumen. Below is a

33

brief description of the mechanisms thought to be used by phagolysosomes for the degradation

of various lipid species.

i) Sphingolipids. Saposins extract glycosphingolipids from bilayers, exposing them to soluble

lipases such as glycosidases (Fig. 1.5c). Similarly, the GM2-activator protein (GM2-AP)

presents GM2 and GM1 gangliosides to water-soluble hydrolases (Fig. 1.5c) (Kolter and

Sandhoff, 2010). Sphingomyelinase, in contrast, can hydrolyze membrane-resident

sphingomyelin in the absence of lipid-binding proteins (Fig. 1.5c) (Linke et al., 2001). Both acid

sphingomyelinase and prosaposins (the precursor proteins that give rise to the saposins) are

delivered to the phagosome from the Golgi in a sortilin-dependent manner (Wähe et al., 2010).

The degradation of sphingolipids converges on ceramide, which is converted into sphingosine

by ceramidase (Fig. 1.5c). Some evidence suggests that the Niemann-Pick disease proteins C-

1 and -2 (NPC1 and NPC2), which are discussed below, may be involved in the export of

sphingosine from lysosomes and likely also from phagolysosomes (Fig. 1.5c). However, the

endo-lysosomal accumulation of sphingolipids seen in Niemann-Pick type C disease patients

(Lloyd-Evans et al., 2008) may be an indirect consequence of excessive cholesterol

accumulation.

ii) Cholesterol. Esterified cholesterol contained in phagocytic targets such as apoptotic bodies,

is de-esterified by the lysosomal acid lipase (Fig. 1.5c). This hydrolase has been detected in

phagosomes through proteomics studies (Trost et al., 2009). The unesterified cholesterol must

be exported from the compartment to prevent its accumulation. Cholesterol transport across the

phagosomal membrane is likely to involve the NPC proteins. NPC2 is a soluble protein that

binds non-esterified cholesterol with high affinity (Friedland et al., 2003). Strong evidence

supports the idea that NPC2, which is found in late compartments of the endocytic and

phagocytic pathway, ferries cholesterol from the lysosomal lumen to NPC1 (Fig. 1.5c), an

integral lysosomal membrane protein (Infante et al., 2008). While the mechanism whereby

34

cholesterol enters the cytoplasm is unknown, it is likely to involve NPC1 (Fig. 1.5). Indeed,

Niemann-Pick disease type C is due to mutations in the genes encoding for NPC1 or NPC2.

The disease is characterized by endo-lysosomal accumulation of cholesterol and diverse

sphingolipids, such as GM2 and GM3 gangliosides, and sphingomyelin (Liscum, 2000). Normal

cholesterol exit from lysosomes and/or phagosomes is essential for maturation to proceed to

completion. In cells from Niemann-Pick disease type C patients phagosomes acquired Rab7,

but the GTPase could not be activated and fusion with lysosomes was impaired (Huynh et al.,

2008). Interestingly, ABC transporters similar to those that transport cholesterol at the PM have

been identified in late compartments (van der Kant et al., 2013b), but whether they play a role in

the export of cholesterol from the phagosome is unclear.

iii) Glycerophospholipids. The lysosomal phospholipase A2 (LPLA2) that can cleave

glycerophospholipids is highly expressed in macrophages (Hiraoka et al., 2006; Schneider et

al., 2014) and has been found in phagosomes (Fischer et al., 2001). Lack of LPLA2 in mouse

macrophages causes phospholipid accumulation, generation of inclusion bodies and foam-cell

formation (Hiraoka et al., 2006). Whether and how lysophospholipids generated by the lipase

are transported from the phagosome to the cytosol is currently unknown. However, following

phagocytosis bacterial cardiolipin is hydrolyzed by LPLA2, yielding lysocardiolipin. This product

is exported from the phagosome. In addition to intracellular phospholipases, secreted

phospholipases, such as group IIA phospholipase A2, can be internalized along with phagocytic

cargo. Internalized phospholipases synergize with intraphagosomal ROS production to facilitate

the breakdown of intraluminal phospholipids (Femling et al., 2005).

The mechanisms whereby lipids are processed after traversing the phagolysosomal membrane

are virtually unknown (Fig. 1.5c). Recent work has revealed the existence of membrane contact

sites specialized in inter-organelle lipid transfer. A distinct possibility is that lipids are

transferred via such membrane contact sites (or via vesicular traffic) to lipid-processing

35

organelles like the ER and the Golgi. From these organelles, lipids can be directed to their

destinations, which can include lipid antigen presentation, storage in fat droplets, or utilization

by the phagocyte for renewal of its membranes of for metabolic consumption.

iv) Limiting phagosomal membrane. As described above, the limiting membrane of the

phagosome undergoes drastic transformations during the maturation process. During early and

intermediate stages of maturation components of the phagosomal membrane can be recycled to

the PM, delivered to the TGN or extruded inward to form ILVs intended for degradation.

However, the fate of the limiting membrane of the mature phagolysosome remains poorly

studied. Once its microbicidal and degradative obligations are fulfilled, phagosomes must be

resorbed. Some evidence points at a ‘very late’ maturation stage characterized by mTORC1-

dependent fission of the phagolysosome (Fig. 1.5c) (Krajcovic et al., 2013), analogous to the

reported tubulation of autophagosomes (Yu et al., 2010) and lysosomes (Saric et al., 2016).

Interestingly, membrane tubulation and vesicular traffic have been linked to a crucial

immunological process: antigen presentation (reviewed in Mantegazza et al., 2013). Hence, it is

imaginable that resolution of the phagolysosomal membrane is coupled to antigen presentation

mechanisms. Finally, different lines of evidence point to the existence of post-lysosomal

compartments. These organelles have been proposed to traffic towards the cell periphery, in

the process losing some lysosomal properties (Padh et al., 1993; Johnson et al., 2016). It is

therefore conceivable that during the very late stages of phagolysosomal maturation, specific

membrane components undergo fission and are selectively transported to the cell periphery.

Such centrifugal displacement could be mediated by Arl8B, a GTPase that associates with

kinesin via its effector SKIP (Fig. 1.5) (Rosa-Ferreira and Munro, 2011).

The notion that phagosomes need to undergo resolution has not been given sufficient attention.

As a consequence, the molecular aspects and implications of resolution are virtually

unexplored. I believe they are crucial to the biology of phagocytes.

36

1.6 Phosphoinositides in phagocytosis

The life cycle of phagosomes is governed by membrane remodeling and sorting events.

Changes in lipid composition—highly coordinated in space and time—orchestrate several

essential phenomena during each stage of the life cycle of phagosomes. Specifically,

phosphoinositides are versatile signaling molecules. This is based on the seven different lipid

species that phosphoinositide metabolism yields. In the cellular environment, the seven species

can be dynamically interconverted by the action of lipid kinases and phosphatases. Yet, the key

aspect of these lipids as master regulators of cellular processes is their ability to selectively

recruit effector proteins. Phosphoinositides achieve this feat, by binding—and hereby

recruiting—domains contained within these effectors with exquisite specificity. The pivotal roles

played by phosphoinositides during phagocytosis are accentuated by evidence that shows that

impairment of their metabolism negatively affects the progression of the process at different

stages.

The sections below aim to describe the changes in phosphoinositide levels as phagocytes

switch from a resting state into active mode during phagocytosis. Additionally, I detail the

enzymatic machinery responsible for these localized changes in phosphoinositide levels. Then,

I attempt to describe the physiological implications of phosphoinositides during each stage of

phagocytosis. Finally, I exemplify molecular mechanisms whereby bacterial pathogens interfere

and commandeer phosphoinositide metabolism, compromising the immune response.

37

1.6.1 Phosphoinositides during phagosome formation

Phosphatidylinositol 4,5-bisphosphate

The primary location of PtdIns(4,5)P2 in resting phagocytes is the inner leaflet of the

plasmalemma, where it encompasses approximately 2 mol % of the phospholipid content

(McLaughlin and Murray, 2005). PtdIns(4,5)P2 is primarily generated by type I

phosphatidylinositol phosphate kinases (PIPKI or PIP5K), which phosphorylate PtdIns4P at the

D5 position of the inositol moiety. Although to a lesser extent, PtdIns(4,5)P2 is also synthesized

through the phosphorylation of PtdIns5P at the D4 position by type II PIPK (Bohdanowicz and

Grinstein, 2013). Dephosphorylation of PtdIns(3,4,5)P3 by phosphatase and tensin homolog

(PTEN) also represents a source for the resting pool of PtdIns(4,5)P2 (Mondal et al., 2011), but

the relative contribution of this pathway is not well established.

The PIP5K family is comprised of three isoforms (PIP5Kα, β and γ). Though PIP5K isoforms

associate with the Golgi complex and tubular lysosomes (Brown et al., 2001; Hammond et al.,

2009), they predominantly distribute to the PM, the most negatively charged compartment in the

cell, through a polycationic region located on their surface (Fairn et al., 2009). Rho (Tolias et

al., 1995; Weernink et al., 2004) and Arf (Brown et al., 2001) GTPases positively regulate PIP5K

activity, thereby integrating diverse processes—such as cytoskeletal remodeling and membrane

traffic—with PtdIns(4,5)P2 synthesis. The relationship between PIP5K and small GTPases

appears to be reciprocal; not only do Rho GTPases control PIP5K activation (Weernink et al.,

2004), but actin polymerization driven by these GTPases is in turn dependent on the activity of

PIP5K (Tolias et al., 1998, 2000). PIP5K is also activated by Arf6, which colocalizes with PIP5K

at sites of PtdIns(4,5)P2 enrichment, including membrane ruffles (Honda et al., 1999; Brown et

al., 2001). Of note, Arf6-mediated activation of PIP5K is strictly dependent on the presence of

anionic phospholipids, such as PtdOH (Honda et al., 1999). PtdIns(4,5)P2 and PtdOH likely

engage in a positive feedback loop, as PtdIns(4,5)P2 is a cofactor necessary for optimal activity

38

of phospholipase D (PLD) (Divecha et al., 2000), an enzyme that forms PtdOH by hydrolyzing

PtdCho.

Counter-balancing the anabolic pathways described above are at least three independent

mechanisms that break down PtdIns(4,5)P2. Class I PI3K catalyzes the phosphorylation of

PtdIns(4,5)P2, converting it to PtdIns(3,4,5)P3. A wide number 5-phosphatases, including OCRL

(Mehta et al., 2014), INPP5B (Bohdanowicz et al., 2012b) and the synaptojanins (Voronov et al.,

2008), promote its degradation to PtdIns4P. PLCγ hydrolyzes the PtdIns(4,5)P2 head-group,

thus forming plasmalemmal DAG and releasing Ins(1,4,5)P3 into the cytosol.

PtdIns(4,5)P2 exerts many important functions at the membrane. It binds proteins with polybasic

motifs or with PH, FERM or ENTH domains (McLaughlin and Murray, 2005), and can also be

converted to second messengers, such as PtdIns(3,4,5)P3, DAG and Ins(1,4,5)P3, which play

crucial roles in the formation and maturation of phagosomes. PtdIns(4,5)P2 and its metabolites

control a remarkable number of events in phagocytosis. These include rearrangement of the

actin cytoskeleton (Rohatgi et al., 2000a) and the accompanying changes in phagocytic

receptor mobility (Jaumouillé and Grinstein, 2011), integrin activation (Martel et al., 2001),

membrane traffic (Simonsen et al., 2001; Di Paolo et al., 2004), plasmalemmal-cytoskeletal

linkages (Hamada et al., 2000), and ion channel activity (Suh and Hille, 2005). Notably, each of

these events takes place in a discrete cellular location and at a particular time during the course

of phagocytosis. The latter raises a recurrent question: how can a single phosphoinositide that

is present throughout the PM orchestrate such spatiotemporally restricted phenomena? Part of

the answer is that PtdIns(4,5)P2 levels change only locally during phagocytosis (Kutateladze,

2010). Microscopy-based determinations in single macrophages expressing a fluorescent

chimera of the PH domain of PLCδ [a PtdIns(4,5)P2 reporter] revealed biphasic changes at the

membranes of forming phagosomes (Botelho et al., 2000). In contrast, PtdIns(4,5)P2 levels

remain steady during the course of phagocytosis in the unengaged aspects of the plasmalemma

39

(Botelho et al., 2000). Fig. 1.6 illustrates that, while there is a noticeable accumulation of

PtdIns(4,5)P2 in emerging pseudopods during the early stages of phagosome formation, its

concentration drops at the base of the phagocytic cup as pseudopodia extend. Following

phagosome sealing and severing, phagosomal PtdIns(4,5)P2 decreases precipitously and is no

longer detectable by fluorescence microscopy (Botelho et al., 2000). In principle, the localized

initial accumulation of PtdIns(4,5)P2 at the pseudopods could result from an upsurge in

synthesis, a decline in consumption, or both. Two lines of indirect evidence point towards an

increase in synthesis. First, activation of PLD has been detected during phagocytosis (Kusner

et al., 1999; Lee et al., 2002; Iyer et al., 2004), this enzyme promotes the recruitment and

activation of PIP5K (Divecha et al., 2000). The simultaneous activation of Arf6 and PLD are

likely to stimulate PIP5K, thereby promoting PtdIns(4,5)P2 formation. Second, all three PIP5K

isoforms transiently accumulate at the phagosomal membrane during the early phase of

phagocytosis, and detach during the later stages (Fairn et al., 2009), correlating with the

biphasic nature of phagosomal PtdIns(4,5)P2 levels. The top panel of Fig. 1.7 illustrates the

signaling pathways leading to PIP5K activation and the consequent formation of phagosomal

PtdIns(4,5)P2.

In turn, as shown in Fig. 1.7b the disappearance of PtdIns(4,5)P2 from the base of phagocytic

cups and eventually from the membrane of sealed phagosomes is mediated by a combination of

kinases, phosphatases and lipases. Specifically, at later stages of phagosome formation PI3K

promotes the phosphorylation of PtdIns(4,5)P2 into PtdIns(3,4,5)P3, which also serves as a

signal for the recruitment of PLCγ (Falasca et al., 1998). Indeed, having being recruited to sites

of PtdIns(3,4,5)P3 production, PLCγ becomes largely responsible for the disappearance of

PtdIns(4,5)P2 (Azzoni et al., 1992; Liao et al., 1992). The inositol 5-phosphatases OCRL and

INPP5B also contribute by converting PtdIns(4,5)P2 into PtdIns4P.

40

Figure 1.6. Distribution of PtdIns(4,5)P2 during phagocytosis. The phagocytic response has been broken down into four conceptual stages: particle recognition (a); extension of pseudopodia (b); membrane fusion/particle internalization (c); and formation of an early phagosome (d). Left) Time-lapse

41

fluorescence images of an RAW 264.7 macrophage expressing PLCδ(PH)-GFP, a PtdIns(4,5)P2-specific fluorescent biosensor. Images were acquired by confocal microscopy immediately after macrophages were challenged with IgG-opsonized targets. PtdIns(4,5)P2 is present in the inner monolayer of the plasmalemma, and its concentration increases locally at sites of particle binding and at forming pseudopods. However, the phosphoinositide is depleted from the base of phagocytic cup as pseudopodia progress and becomes undetectable at the phagosomal membrane following scission. Phagocytic particles are denoted with a star. Scale bar, 5 µm. Right) Schematic representation of the local changes in PtdIns(4,5)P2 concentration during phagocytosis, corresponding to the experimental data obtained with the PLCδ (PH)-GFP probe. The color code of the membrane is indicative of the relative abundance of PtdIns(4,5)P2, ranging from grey (lowest) through light green (intermediate), to dark green (highest). Fc receptors are shown in orange; the opsonin, IgG, in blue and the phagocytic target in violet.

In addition to the catabolic pathways described above, detachment of PIP5K from internalized

phagosomes facilitates the exclusion of PtdIns(4,5)P2 from these compartments. The

dissociation of PIP5K has been attributed to a localized and acute drop in phagosomal surface

charge (Yeung et al., 2006). The release of PIP5K from sealed phagosomal membranes

terminates PtdIns(4,5)P2 synthesis and further promotes the disappearance of this lipid at a site

where degradation is already ongoing.

In phagocytes, as in many other cellular systems, sites of PtdIns(4,5)P2 formation serve as

signaling platforms that trigger robust actin polymerization. PtdIns(4,5)P2 promotes the

activation of a number of actin-regulatory proteins that are responsible for filament assembly,

while inhibiting those in charge of disassembly (Saarikangas et al., 2010) (Fig. 1.7a). Proteins

that directly bind to actin and dictate the equilibrium between its monomeric and filamentous

form include profilin (Chaudhary et al., 1998), cofilin (Gorbatyuk et al., 2006), gelsolin (Janmey

and Stossel, 1987) and capping protein (Cooper and Sept, 2008). In addition to increasing the

number of barbed ends, PtdIns(4,5)P2 induces de novo actin nucleation by activating NPFs

42

Figure 1.7. Functional implications of PtdIns(4,5)P2 metabolism for phagocytosis. a) Pathways leading to PtdIns(4,5)P2 synthesis (main panel) and consequent stabilization of F-actin networks (lower panels) at the phagocytic cup. Membranes are colored-coded as in Figure 1. Rac1 and Arf6 activate PIP5K in a PtdOH-dependent fashion. PtdOH can be synthesized through the phosphorylation of DAG by DGK or through hydrolysis of PtdCho by PLD. PtdOH recruits Rac GEFs (e.g. Tiam1) to phagocyte membranes, where Rac is activated. In turn, PIP5K associates with the plasma membrane through a

43

positively charged surface, where it catalyzes the conversion of PtdIns4P to PtdIns(4,5)P2. PtdIns(4,5)P2

mediates linkage of actin networks to integral plasmalemmal proteins through intermediary ERM proteins, as well as actin polymerization by nucleation-promoting factors such as WASp. In addition, actin-biding proteins that antagonize filament formation, such as capping protein and the severing factor cofilin, are inhibited by PtdIns(4,5)P2. b) Depletion of PtdIns(4,5)P2 from the base of the cup leads to actin filament removal. PtdIns(4,5)P2 is converted by kinases (PI3K), phosphatases (OCRL) and phospholipases (PLCg). Disappearance of the filamentous actin barrier facilitates the delivery of membranes from endolysosomal compartments to the phagocytic cup, releasing membrane tension. c) PLCg-mediated PtdIns(4,5)P2 hydrolysis results in the formation of the bioactive molecules DAG and Ins(1,4,5)P3 (IP3). Regulatory subunits of the NADPH oxidase are activated by PKC, which is recruited to phagosomal membranes by DAG. Release of Ca2+ from intracellular stores is promoted by IP3.

(Miki et al., 1996). Lastly, ezrin/moesin/radixin (ERM), which directly link the cytoskeleton to the

plasmalemma, are also well established PtdIns(4,5)P2 effectors (Bretscher et al., 2002).

Because of these effects, the local increase in PtdIns(4,5)P2 synthesis that occurs upon

engagement of receptors causes a large-scale reorganization of the actin cytoskeleton, driving

the extension of pseudopodia around the surface of phagocytic targets (Coppolino et al., 2002).

This claim is supported by studies in which expression of a kinase-dead PIP5K that impairs the

formation of phagosomal PtdIns(4,5)P2 precluded accumulation of F-actin in nascent phagocytic

cups, and depressed the phagocytic capacity of the cells (Coppolino et al., 2002).

Although PIP5Kα, β and γ are all recruited to nascent phagosomes, different isoforms have

been reported to mediate distinct, non-redundant roles during phagocytosis (Mao et al., 2009).

PIP5Kα activity has been implicated in the activation of the nucleation promoting factor WASp

(Mao et al., 2009), which catalyzes actin polymerization and pseudopod extension (Park and

Cox, 2009). In contrast, PIP5Kγ seems to control the mobility of phagocytic receptors in the

plane of the membrane, presumably by modulating the density of cortical actin. Subsequently,

PIP5Kα catalyzes the emission of pseudopodia by promoting WASp activity (Mao et al., 2009).

To reconcile the fact that these isoforms have divergent effects, it has been postulated that

PIP5Kγ is subjected to post-translational control by Syk, thereby restricting its activity to a

particular region and time (Mao et al., 2009).

44

While expansion of the actin skeleton and its anchorage to the plasmalemma drive formation of

pseudopodia at early phases of phagocytosis, phagosome scission is accompanied by the

disappearance of actin from the base of the phagocytic cup (Scott et al., 2005). Indeed, actin

clearance is a requirement for completion of phagocytosis, especially of large phagocytic targets

(Cox et al., 1999a; Beemiller et al., 2010). Abortive phagocytic cups develop when the

disintegration of the phagosomal actin meshwork is prevented either by expressing

constitutively-active Rho GTPases (Beemiller et al., 2010) or by inhibiting PI3K (Araki et al.,

1996; Cox et al., 1999a). The role of PtdIns(3,4,5)P3 in actin disassembly is discussed in detail

below.

The catabolism of PtdIns(4,5)P2 coincides in space and time with the breakdown of actin (Scott

et al., 2005). The loss of phagosomal actin occurs asymmetrically after phagosomal sealing,

with depolymerization arising initially at the base of the phagocytic cup, strongly resembling the

pattern of PtdIns(4,5)P2 disappearance. Of note, dismantling of actin at the base of the cup and

particle internalization are blocked if high PtdIns(4,5)P2 levels are sustained by promoting

PIP5K-mediated synthesis or by inhibiting PLCγ-driven degradation (Scott et al., 2005). It thus

appears that loss of PtdIns(4,5)P2 causes actin disassembly, which is in turn required for

completion of phagocytosis.

In addition to the consequences that PtdIns(4,5)P2 metabolism has on cytoskeletal dynamics,

breakdown of the inositide to secondary metabolites also has important ramifications (Fig. 1.7c).

PLCγ-mediated hydrolysis of PtdIns(4,5)P2 leads to the formation of DAG and Ins(1,4,5)P3. The

kinetics and spatial distribution of DAG liberation during phagocytosis have been measured with

a genetically encoded fluorescent chimera of the C1 domain of PKCδ (Botelho et al., 2000),

which selectively associates with DAG (Oancea et al., 1998). Consistent with the role of PLCγ

45

in phagocytosis, appearance of DAG coincides in space and time with the disappearance of

PtdIns(4,5)P2. Unexpectedly, though neither DAG nor Ins(1,4,5)P3 are essential for particle

engulfment, inhibition of PLCγ blocks the phagocytic response (Botelho et al., 2000; Scott et al.,

2005). It thus appears that disappearance of PtdIns(4,5)P2, rather than the formation of its

metabolites, may be essential for completion of phagocytosis.

While DAG and Ins(1,4,5)P3 are not required for particle internalization, recruitment of

conventional and novel PKC isoforms by DAG, as well as Ca2+ mobilization by Ins(1,4,5)P3, play

significant roles in other stages of phagocytosis (Bengtsson et al., 1993; Ueyama et al., 2004;

Nunes et al., 2012; Schlam et al., 2013). Recruitment and activation of PKC by DAG during

phagocytosis affect the elimination of internalized pathogens, as PKC phosphorylates and

activates p47phox, a regulatory subunit of the NADPH oxidase (NOX) (He et al., 2004; Cheng et

al., 2007). Underscoring the significance of DAG formation is the observation that NOX

activation is precluded if individual phagosomes fail to reach a critical DAG concentration

(Schlam et al., 2013).

In addition to stimulating PKC, DAG represents a source of PtdOH, which is synthesized by

DAG kinases (Bohdanowicz et al., 2013). PtdOH biosynthesis is particularly important for

professional phagocytes; in contrast to non-phagocytic cells, macrophages and dendritic cells

display elevated levels of PtdOH at their plasma membranes (Bohdanowicz et al., 2013).

Preceding phagocytosis and macropinocytosis, phagocytes continuously survey their

environment in the lookout for foreign particles or soluble antigens. This incessant ruffling is

strictly dependent on the constitutive conversion of DAG to PtdOH, which is in turn responsible

for the recruitment of Rac1 GEFs, including TIAM1, to the plasmalemma (Bohdanowicz et al.,

2013). Activation of Rac1 by TIAM1 then promotes nucleation of actin filaments and the

extension of membrane protrusions that facilitate capture of phagocytic targets.

46

As shown in Figure 1.7c, PtdIns(4,5)P2 hydrolysis is accompanied by the release of Ins(1,4,5)P3

into the cytosol and its diffusion to the ER, where it induces Ca2+ release by binding to the

Ins(1,4,5)P3 receptor (Nunes et al., 2012). Depletion of Ca2+ from intracellular stores is sensed

by STIM1, a transmembrane ER-resident protein. A recent study demonstrated that, upon

depletion of calcium from the ER lumen, STIM1 recruits ER cisternae to nascent phagosomes

and promotes opening of store-operated calcium entry channels present at the phagosomal

membrane (Nunes et al., 2012). Earlier studies implicated an increase in free cytosolic Ca2+ in

the fusion of secondary granules with the PM of neutrophils (Jaconi et al., 1990). Ca2+ influx

may play a similar role in the focal delivery of endomembranes to sites of phagocytosis.

Comparatively little is known about the regulation of phosphatases during phagocytosis and

their contribution to PtdIns(4,5)P2 removal. Nonetheless, evidence pointing to a role for OCRL

and INPP5B has begun to emerge (Bohdanowicz et al., 2012a). These inositol 5-phosphatases

are Rab5 effectors, and associate with nascent phagosomes through an adaptor protein called

APPL1, also a Rab5 effector (Bohdanowicz et al., 2012a). Silencing of either APPL1 or Rab5

prolonged the accumulation of both PtdIns(4,5)P2 and filamentous actin on phagosomal

membranes (Bohdanowicz et al., 2012a). These observations are consistent with previous

studies in Dictyostelium, where inactivation of Dd5P4 (a homolog of OCRL), resulted in

phagocytic impairments (Loovers et al., 2007). Remarkably, although phagocytic cups did form

in Dd5P4-null cells, these did not manage to seal and remained at an abortive stage (Loovers et

al., 2007). Recent experimental findings have implicated a trimeric complex consisting of Bcl10

and the clathrin adaptors AP1 and EpsinR in ferrying OCRL to nascent phagosomes (Marion et

al., 2012). In these studies, depletion of Bcl10 resulted in the formation of unproductive

phagocytic cups, rich in PtdIns(4,5)P2, Cdc42 and F-actin (Marion et al., 2012).

It is evident that PtdIns(4,5)P2 metabolism has profound implications for the formation and

maturation of phagosomes. Thus, it is not surprising that microbes often subvert PtdIns(4,5)P2

47

signaling in order to colonize their host. In an evolutionary arms race, a number of intracellular

microbes have developed the ability to hijack the polymerization of actin belonging to the host

cells to gain entry and move within them (Frischknecht and Way, 2001). Active modulation of

plasmalemmal PtdIns(4,5)P2 homeostasis by secreted effectors is one of the strategies used by

invasive bacteria to co-opt the host actin-regulatory machinery. This is particularly important

because most of the tension of the PM (≈75%) is thought to result from its coupling to cortical

actin (Sheetz, 2001), which is in turn dependent on PtdIns(4,5)P2. By promoting breakdown of

PtdIns(4,5)P2 intracellular pathogens weaken cytoskeletal support and reduce membrane

rigidity, facilitating entry (Terebiznik et al., 2002).

Salmonella spp., which are facultative intracellular pathogens, have evolved a number of

virulence mechanisms that enable their entry into host cells (Fu and Galán, 1999). These

bacteria deliver effector proteins into host cells via a type III secretion system (T3SS), a

specialized needle-like molecular machine (Fu and Galán, 1999), causing massive

reorganization of the host’s actin network. In conjunction with SopE and SopE2, two effectors

that act as mimics of GEFs for Rho GTPases, Salmonella translocates a phosphoinositide

phosphatase, SopB, that rapidly eliminates PtdIns(4,5)P2 from the invaginating regions of

membrane ruffles (Terebiznik et al., 2002). The disappearance of PtdIns(4,5)P2 facilitates

membrane deformation, enabling both extension of ruffles as well as the scission of the

Salmonella-containing vacuoles (SCV). Remarkably, SopB-mediated invasion is not restricted

to the subversion of cytoskeletal dynamics; this protein also diverts phagosome maturation to a

non-lytic compartment by reducing the surface charge of the SCV (Bakowski et al., 2010). By

reducing the levels of anionic phospholipids, SopB precludes association of specific Rab

GTPases that carry polycationic tails with the SCV. These proteins rely on electrostatic forces

for their localization, and their displacement interferes with proper endocytic traffic and

phagolysosome formation.

48

Phosphatidylinositol 3,4,5-trisphosphate

Like other 3-polyphosphoinositides, PtdIns(3,4,5)P3 levels are scarce in unstimulated cells.

While its levels are minute at rest (less than 0.2% of all inositol-containing lipids) (Rameh and

Cantley, 1999), PtdIns(3,4,5)P3 is quickly generated upon engagement of immune receptors.

The metabolism of PtdIns(3,4,5)P3 is strictly and dynamically regulated, and in general

restricted to the cytosolic leaflet of the plasmalemma (Palmieri et al., 2010).

Generation of PtdIns(3,4,5)P3 occurs mainly via phosphorylation of PtdIns(4,5)P2 by the family

of class I PI3K, which localize to the PM and use PtdIns(4,5)P2 as a substrate for

PtdIns(3,4,5)P3 biosynthesis (Botelho et al., 2000). The class I PI3K holoenzymes are

comprised of a regulatory subunit (either p85 or p101) and a catalytic p110 subunit

(Vanhaesebroeck and Waterfield, 1999). While the p85 regulatory subunit acts downstream of

receptor tyrosine kinases, the p101 subunit responds to GPCRs. The breakdown of

PtdIns(3,4,5)P3 occurs mainly through the action of 3- and 5-phosphatases; PTEN

dephosphorylates PtdIns(3,4,5)P3 at the D3 position, generating PtdIns(4,5)P2 (Maehama and

Dixon, 1998), while SHIP hydrolyzes the D5 position, producing PtdIns(3,4)P2 (McCrea and De

Camilli, 2009).

The spatiotemporal dynamics of PtdIns(3,4,5)P3 synthesis during phagocytosis mirror those of

PtdIns(4,5)P2 disappearance, consistent with a role for class I PI3K in mediating the conversion

of PtdIns(4,5)P2 to PtdIns(3,4,5)P3. Indeed, PI3K is recruited to and activated at sites of

phagocytosis (Marshall et al., 2001); upon particle engagement, tyrosine kinases recruit p85,

the regulatory subunit of class I PI3K, initiating PtdIns(3,4,5)P3 formation (Kwiatkowska and

Sobota, 1999). Synthesis of phagosomal PtdIns(3,4,5)P3 is detectable shortly after phagocytic

targets are engaged, and the phosphoinositide continues to accumulate as the phagocytic cup

progresses. While PtdIns(3,4,5)P3 is still detectable after sealing, its presence in the

phagosomal compartment is short-lived and its concentration declines sharply within 1-2

49

minutes of sealing. Notably, SHIP accumulates at the phagosomal membrane (Marshall et al.,

2001), where it likely promotes the breakdown of PtdIns(3,4,5)P3 to PtdIns(3,4)P2 (Kwiatkowska

and Sobota, 1999; Marshall et al., 2001; Kamen et al., 2007). Fig. 1.8 presents a diagrammatic

representation of the spatiotemporal dynamics of PtdIns(3,4,5)P3 during phagocytosis.

PtdIns(3,4,5)P3 plays a critical and pleiotropic role during phagocytosis. Accordingly, a

profound impairment of phagocytosis is observed in macrophages treated with PI3K inhibitors.

Tellingly, treatment with the PI3K inhibitors wortmannin or LY294002 results in the formation of

abortive cups that do not extend fully around the particle’s circumference. However, the

polymerization of actin and the initial extension of pseudopodia persist in inhibitor treated cells.

Thus, PI3K is dispensable for the initial stages of actin polymerization, but is necessary for later

stages of pseudopodial progression, and perhaps even for phagosome sealing. Interestingly,

the dependency of phagocytosis on PI3K seems to be a size-dependent phenomenon, as the

uptake of large particles is much more affected by PI3K inhibitors than that of small ones (Cox

et al., 1999a). It is worth emphasizing that the unproductive phagocytic cups that form when

PI3K is inhibited stall at a stage where filamentous actin is richly accumulated at the base of the

cup. The latter observation suggests that PtdIns(3,4,5)P3 may be necessary for mediating actin

breakdown, perhaps by allowing for actin, or determinants of its polymerization, to be recycled

to the tips of advancing pseudopods. Orchestration of actin clearance by PI3K likely arises as

a combined effect of PtdIns(4,5)P2 removal and the inactivation of Rho GTPases. Consistent

with this notion, it has been suggested that PI3K negatively regulates Cdc42 at later stages of

phagocytosis (Beemiller et al., 2010).

Like other phosphoinositides, PtdIns(3,4,5)P3 orchestrates its many cellular functions by

recruiting effectors that carry domains that specifically recognize its head-group. Several of

these effectors carry PH domains, including myosin X, an unconventional motor protein that has

been implicated in pseudopodial extension and phagosome closure (Cox et al., 2002).

50

Figure 1.8. Distribution of PtdIns(3,4,5)P3 during phagocytosis. The phagocytic response is broken down into the same stages defined in Figure 1.6. Left) Time-lapse fluorescence images of a RAW 264.7 macrophage expressing Akt(PH)-GFP, a fluorescent probe that detects PtdIns(3,4,5)P3 [and also

51

PtdIns(3,4)P2]. While virtually absent from the bulk plasmalemma, engagement of phagocytic receptors triggers a transient yet marked accumulation of PtdIns(3,4,5)P3 in the membrane of the nascent phagosome. This increase persists until phagosome sealing, then PtdIns(3,4,5)P3 is depleted from the phagosomal membrane shortly (1-2 min) after internalization. Phagocytic particles are denoted with a star. Scale bar, 5 µm. Right) Schematic representation of the local changes in PtdIns(3,4,5)P3

concentration at sites of phagocytosis, corresponding to the experimental data obtained with Akt(PH)-GFP. The color code of the membrane is indicative of the relative abundance of PtdIns(3,4,5)P3, ranging from grey (lowest), through light red (intermediate), to dark red (highest). Other details as in Figure 1.6.

Treatment with wortmannin blocks myosin X enrichment at the phagocytic cup, and the

expression of a truncated form of this motor reduces the ability of macrophages to carry out

FcγR-dependent phagocytosis. In this regard, it is interesting that inhibition of myosin X activity

prevented spreading but not adhesion of macrophages on IgG-coated substrates, and inhibited

phagocytosis of large particles (Cox et al., 2002). Thus, it has been suggested that the

dependency of phagocytosis of large particles on PI3K activity may be attributable to the

recruitment of myosin X. Fig. 1.9 illustrates the practical implications of PtdIns(3,4,5)P3

metabolism for phagosome formation.

PtdIns(3,4,5)P3 signaling is also often hijacked by intracellular pathogens as part of their

colonizing strategy. Enteropathogenic Escherichia coli (EPEC) invades the intestinal epithelium

by inducing the formation of F-actin-rich pedestals by a process that relies on the subversion of

PtdIns(3,4,5)P3 homeostasis (Sason et al., 2009). These actin-driven structures facilitate

colonization and increase pathogenicity by allowing the bacteria to adhere tightly to intestinal

surfaces (Frischknecht and Way 2001). Using a T3SS, EPEC inject a protein called

translocated intimin receptor (Tir). The extracellular region of Tir acts as a receptor for intimin,

a bacterial adhesin that operates as an ‘attach and efface’ virulence factor. Engagement of

intimin by Tir leads to the clustering of the translocated bacterial receptors on the plane of the

host cell membrane, which in turn recruits PI3K and triggers a cascade of actin rearrangement

events that require PtdIns(3,4,5)P3 (Sason et al., 2009).

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Figure 1.9. Functional implications of PtdIns(3,4,5)P3 metabolism for phagocytosis. a) Signal transduction pathways leading to PI3KI activation at the nascent phagosome. Engagement of FcγR by

53

an IgG-coated target triggers receptor clustering in the plane of the membrane, promoting phosphorylation on ITAM motifs by Src family kinases (SFK). Doubly-phosphorylated ITAMs are sensed by a tandem SH2 domain on the non-receptor tyrosine kinase Syk, which binds to and directly activates the adaptor LAT (not illustrated). LAT stimulates docking of additional, proteins, such as PLCg and Grb2. The former catalyzes PtdIns(4,5)P2 hydrolysis, while the latter acts as an adaptor protein for Gab2. Once recruited to active phagocytic receptors, Gab2 is phosphorylated by Syk and subsequently induces recruitment of p85, the regulatory subunit of PI3KI. The class I PI3K holoenzyme then phosphorylates PtdIns(4,5)P2, generating PtdIns(3,4,5)P3. Note that Gab2 is stabilized at the phagosomal membrane by PtdIns(3,4,5)P3 produced by PI3KI, thereby amplifying the PtdIns(3,4,5)P3 signal. b) Orchestration of pseudopodia progression by direct PtdIns(3,4,5)P3 effectors. PLCγ is recruited to the phagocytic cup by a PtdIns(3,4,5)P3-interacting PH domain, promoting PtdIns(4,5)P2 breakdown. Disappearance of PtdIns(4,5)P2 results in the removal of F-actin from the base of the cup. Termination of polymerization is reinforced by the PtdIns(3,4,5)P3-mediated recruitment of Rho GAPs and the consequent deactivation of Rho GTPases. Myosin motors (e.g. myosin X) also translocate to the cup in a PtdIns(3,4,5)P3 dependent manner, where they facilitate phagosome sealing by exerting contractile forces.

1.6.2 Phosphoinositides during phagosome maturation

Phosphatidylinositol 3-phosphate

Though its cellular concentration is comparatively low, PtdIns3P is critically involved in the

maturation of phagosomes. In mammalian cells, PtdIns3P is found mainly at the cytoplasmic

leaflet of early endosomes and in ILVs of multivesicular bodies (Gillooly et al., 2000). The

predominant source of this inositide is class III PI3K (Vps34), which phosphorylates the D3

position of PtdIns (Stephens et al., 1994; Backer, 2008). Vps34 localizes to early endosomes

(Yan and Backer, 2007), and its inhibition by wortmannin (Stephens et al., 1994) or by specific

anti-Vps34 antibodies (Vieira et al., 2001) quickly eliminates PtdIns3P from these

compartments. Though quantitatively less predominant, other sources of PtdIns3P also exist:

phosphorylation of PtdIns by class II PI3K (MacDougall et al., 1995) and dephosphorylation of

bisphosphorylated species by inositol polyphosphate phosphatases (Ferron and Vacher, 2006).

In principle, three possible mechanisms could account for disappearance of PtdIns3P:

phosphorylation, dephosphorylation and hydrolysis. The precise relative contribution of these

pathways is not clear. However, enzymes that could potentially carry each of these functions

have been identified. PIKfyve eliminates PtdIns3P by phosphorylating its D5 position,

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generating PtdIns(3,5)P2 (Burd and Emr, 1998). Conversely, PtdIns3P can be broken down by

the 3-phosphatase MTM1, a member of the myotubularin family. In addition to its hydrolytic

activity, MTM1 directly interacts with Vps34 and competitively displaces it from endosomal

membranes, thereby preventing it from engaging Rab5 or Rab7 (Yan and Backer, 2007).

Lastly, PtdIns3P could be removed by lysosomal phospholipases, which gain access to the

inositide as ILVs form upon ESCRT-mediated invagination of the limiting membrane (Ching et

al., 1999).

When phagocytes encounter a target, PtdIns3P is initially absent from pseudopodia and the

neighboring (unengaged) plasmalemma. However, sealing of the phagosome and its

internalization is followed by a striking yet transient accumulation of the phosphoinositide, which

lasts for about 10 min and coincides with the centripetal movement of the phagosomal vacuole

(Vieira et al., 2001). The disappearance of phagosomal PtdIns3P is predicted to involve a

combination of phosphorylating and hydrolytic reactions, as well as inward budding. Fig. 1.10

depicts the dynamic changes in the subcellular distribution of PtdIns3P throughout the different

stages of phagocytosis, as measured by a fluorescent biosensor that specifically recognizes this

phosphoinositide.

Its spatiotemporal dynamics suggest that PtdIns3P is dispensable for pseudopod formation but

that its function is related to phagosomes maturation. This notion has been amply validated by

a number of studies where phagosome maturation was precluded by pharmacological inhibition

of PI3K. In these experiments phagocytes were treated with wortmannin, an inhibitor of both

class I and III PI3K, prior to being challenged with small (3 µm) particles (Vieira et al., 2001).

Small particles were utilized in the study because, as discussed above, inhibition of class I PI3K

impairs phagocytosis of large particles, while internalization of small particles is only slightly

affected (Cox et al., 1999a). Under these conditions, phagosomes formed but did not acquire

PtdIns3P. More importantly, these phagosomes arrested at an immature stage that had a

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Figure 1.10. Distribution of PtdIns3P during phagocytosis. Four conceptual stages are shown: particle recognition (a); extension of pseudopodia (b); early phagosome (c); and late phagosome (d). Left) Time-lapse fluorescence microscopy images of a RAW 264.7 macrophage expressing

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EEA1(2XFYVE)-GFP, a PtdIns3P biosensor. PtdIns3P, which localizes primarily to early endosomal compartments, is undetectable at the plasmalemma. PtdIns3P is also absent from phagosomal membranes during particle recognition and pseudopod extension, but accumulates noticeably in the early maturing phagosome (between 1 and ≈10 min after sealing). However, PtdIns3P is lost as the phagosomal vacuole matures into a late phagosome. Phagocytic particles are denoted by a star. Scale bar, 5 µm. Right) Schematic representation of the local changes in PtdIns3P concentration at sites of phagocytosis, corresponding to the experimental data obtained with the EEA1(2XFYVE)-GFP probe. Other details as in Figure 1.6.

markedly reduced content of lysosomal markers such as LAMP-1. Similar results were

obtained when class III PI3K was neutralized by the injection of specific antibodies (Vieira et al.,

2001). Thus, class I and class III PI3K play distinct roles in the phagocytic process; class I

PI3K is responsible for the synthesis of 3-polyphosphoinositides that control pseudopod

extension and sealing, while class III PI3K orchestrates phagosome maturation by catalyzing

PtdIns3P formation in early phagocytic compartments.

Rab5 and Rab7 are critical regulators of membrane traffic and phagolysosome biogenesis.

However, their activation at phagosomal membranes alone is insufficient to drive phagosome

maturation to completion; inhibition of PI3K blocks the progression of phagosomes, even though

the arrested vacuoles retain active Rab7 (Vieira et al., 2003). The functions of Rab5 and 7 in

maturation seem to be heavily dependent on their ability to associate with p150, a myristoylated

Ser/Thr protein kinase and critical binding partner of Vps34 (Murray et al., 2002; Stein et al.,

2003). Both recruitment of Vps34 to membranes and its catalytic activity are augmented by

binding to p150 (Yan and Backer, 2007). Thus, orchestration of membrane traffic by Rab5 and

Rab7 requires Vps34 activation and the consequent accumulation of PtdIns3P in early

phagosomal compartments.

Following its synthesis during the early stages of maturation, PtdIns3P is responsible for

carrying multiple signaling tasks. The phosphoinositide participates in endosome and

phagosome progression, retrieval of membranes to the plasmalemma, sorting of membrane

proteins to the TGN and targeting of cargo for degradation within ILVs. PtdIns3P is also partly

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responsible for the acquisition of bactericidal properties by the phagosome. The many versatile

roles played by PtdIns3P in the course of phagosome maturation are illustrated in Fig. 1.11.

PtdIns3P exerts its effects by recruiting a number of effectors that possess PX or FYVE

domains. Notably, PtdIns3P is the phosphoinositide with the largest collection of specific

binding partners: the human genome encodes 42 PX domain- and 30 FYVE domain-containing

proteins, most of which selectively recognize PtdIns3P (Lemmon, 2003). A prototypical

example is the tethering molecule EEA1, which carries a FYVE domain in its C-terminus that

recognizes PtdIns3P (Simonsen et al., 1998) and binds to early endosomes (He). EEA1 also

recognizes active Rab5 (Mishra et al., 2010). Thus, Rab5 and its downstream target Vps34

synergize to recruit EEA1 to the early phagosomal membrane. EEA1 is crucial for phagosome

maturation, as it mediates fusion with components of the endocytic pathway by interacting with

syntaxins 6 and 13 (Simonsen et al., 1999; Collins et al., 2002); these SNAREs catalyze

membrane fusion during phagocytosis (Collins et al., 2002). For these reasons, neutralization

of EEA1 function by introduction of inhibitory antibodies results in a blockade in phagosome

maturation, much like that observed in wortmannin-treated cells (Fratti et al., 2001a).

In addition to orchestrating fusogenic events between phagosomes and the early endosomal

system, PtdIns3P is central for sorting phagosomal contents to degradative compartments. Fig.

1.11c shows that phagosomal membrane proteins destined for lysosomal destruction undergo

mono- or polyubiquitylation and are subsequently internalized through invagination of the

phagosome limiting membrane, leading to the formation of ILVs. The activated phagocytic

receptor FcγRIIA is one such protein (Lee et al., 2005).

As discussed above, ESCRT proteins are responsible for the generation of ILVs. Most relevant

to this review is Hrs, a subunit of ESCRT-0 that carries a FYVE domain and interacts with

phagosomal PtdIns3P in a highly specific fashion (Vieira et al., 2004).

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Figure 1.11. Functional implications of PtdIns3P metabolism for phagocytosis. a) The tethering protein EEA1 utilizes its FYVE domain to engage PtdIns3P in both phagosomal and endosomal membranes, where it binds to active Rab5. EEA1 also interacts directly with syntaxins 6 and 13, SNARE proteins that facilitate fusion between phagosomes and early endosomes (EE). b) PtdIns3P promotes NADPH oxidase (NOX) activity and ROS production. p40phox, a cytosolic component of NOX, is recruited to the maturing phagosome through a PX domain that recognizes PtdIns3P. p40phox interacts with other

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subunits of the oxidase, stabilizing the complex on the phagosomal membrane and sustaining ROS generation. c) Role of PtdIns3P in ESCRT-mediated sorting of phagosomal membrane proteins. The ESCRT-0 subunit Hrs utilizes a FYVE domain to associate with PtdIns3P-rich membranes, where the complex recognizes ubiquitylated cargo, such as FcgRIIA. ESCRT-0 then triggers the sequential recruitment of ESCRT complexes I, II and III, culminating in the invagination of the limiting membrane. d) PtdIns3P in the retrieval of phagosomal membrane proteins by the retromer complex. Retromer, consisting of a cargo-recognition Vps trimer and a sorting nexin (SNX) dimer, is recruited to maturing phagosomes, where it promotes recycling of membrane proteins aided by actin-driven membrane tubulation. The SNX proteins of retromer carry PX and BAR domains, which they utilize for the recognition of PtdIns3P-rich membranes and the stabilization of membrane curvature within tubules, respectively.

Indeed, inhibition of PtdIns3P synthesis with wortmannin prevents Hrs recruitment to the

phagosome. More importantly, silencing of Hrs arrests maturation at an early stage, with

retention of markers of early (sorting) endosomes on the phagosomal membrane (Vieira et al.,

2004).

Certain phagosomal components, such as acid hydrolase receptors, are not destined for

degradation to the lysosome and instead are retrogradely ferried to the TGN by retromer.

Notably, the SNX subunits of retromer carry a PX domain, which mediates their tethering to

phagosomal PtdIns3P (Cozier et al., 2002). By binding to PtdIns3P through their PX domain

while concomitantly facilitating membrane curvature through their BAR domain, the SNX

subunits of retromer mediate tubule and vesicle formation for the purposes of retrograde

transport (Fig. 1.11d).

Formation of a complex between class III PI3K and the autophagy related protein Beclin-1

appears to be necessary for retromer function. Mutations in Vps30 (the yeast ortholog of

Beclin-1) lead to sorting and maturation defects, as well as decreased PtdIns3P levels (Burda et

al., 2002). The failure to attain suitable PtdIns3P levels precludes association of the SNX1/2-

SNX5/6 dimer with endosomal membranes (Burda et al., 2002). In C. elegans, clearance of

apoptotic corpses is defective when bec-1 (the Beclin-1 ortholog) is mutated, suggesting that

retromer-dependent transport is a component of efferocytosis. Also, deficiencies in retromer

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levels or in its assembly have been linked to phagocytic defects and amyloid-beta removal in

the brain; beclin-1 mutant microglia do not recruit retromer efficiently to nascent phagosomes,

making them incapable of properly recycling receptors such as CD36 to the plasmalemma

(Lucin et al., 2013). Neurodegenerative consequences can ensue due to the depletion of

phagocytic receptors for apoptotic bodies or cell debris.

An important component of the microbicidal arsenal of phagosomes is the NADPH oxidase

(NOX), an electrogenic complex that generates reactive oxygen species (ROS) in the

phagosomal lumen. As shown in Fig 1.4 and Fig. 1.11b, NOX is a multicomponent system

comprised of two membrane proteins (gp91phox and p22phox), three soluble regulatory proteins

(p40phox, p47phox and p67phox) (Dupré-Crochet et al., 2013) and either Rac1 or Rac2 (Ambruso et

al., 2000; Roepstorff et al., 2008). The membrane-associated subunits form a heterodimeric

flavocytochrome that is responsible for the catalytic activity of the complex, generating

superoxide from NADPH and oxygen. While NOX is inactive in unstimulated cells, phagocytic

signals recruit the ternary complex to sites of particle engagement, where the flavocytochrome

is activated (Nunes et al., 2013). Sustained association of the regulatory subunits of the

oxidase with sealed phagosomes is facilitated by p40phox, which carries a PtdIns3P-binding PX

domain (Ueyama et al., 2007, 2008). Indeed, retention of p40phox on the membrane of sealed

phagosomes is prevented when PI3K is inhibited; importantly, sustained stimulation of NOX is

absent when p40phox is knocked out or when its PX domain is mutated (Tian et al., 2008).

Because of its multifunctional role in transforming the phagosome into a microbicidal machine,

PtdIns3P constitutes an attractive target for invasive organisms whose pathogenicity rests on

the ability to prevent phagolysosome biogenesis. A pertinent example is Mycobacterium

tuberculosis, an intracellular bacterium that manages to survive within the protected confines of

the early phagosome by halting its progression (Pethe et al., 2004). This arrest in maturation

has been attributed to the exclusion of PtdIns3P from the limiting membrane of the

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Mycobacterium-containing vacuole (Fratti et al., 2001b, 2003; Vergne et al., 2003). To this end,

the bacterium sheds mannose-capped lipoarabinomannan (ManLAM), a glycosylated PtdIns

and major component of its cell wall that purportedly interferes with Vps34 activity (Vergne et

al., 2003). Mechanistically, it has been proposed that ManLAM blocks a surge in cytosolic Ca2+

that normally accompanies phagocytosis, ostensibly interfering with Ca2+/calmodulin-dependent

activation of class III PI3K (Vergne et al., 2003). In a synergistic manner, M. tuberculosis

secretes SapM, a phosphatase that hydrolyzes PtdIns3P (Flannagan et al., 2012; Puri et al.,

2013). Depletion of PtdIns3P from the mycobacterial phagosome by this combined strategy

prevents the acquisition of critical mediators of vesicular traffic, such as EEA1 and Hrs, thus

aborting phagosome maturation.

Phosphatidylinositol 3,5-bisphosphate

The cellular abundance of PtdIns(3,5)P2 is extremely low. For a sense of scale, it has been

estimated that there are about 100 PtdIns(4,5)P2 molecules and 10 PtdIns3P molecules for

every PtdIns(3,5)P2 molecule (McCartney et al., 2014). PtdIns(3,5)P2 is synthesized via

phosphorylation of PtdIns3P at the D5 position of the inositol ring by PIKfyve. Both

PtdIns(3,5)P2 and PIKfyve preferentially distribute to late endosomes and the lysosomal network

(McCartney et al., 2014). The converse reaction, dephosphorylation of the D5 position in

PtdIns(3,5)P2, is carried by the SAC domain-containing phosphatase FIG4 (McCartney et al.,

2014). The myotubularin (MTM) family of phosphatases catalyze an alternative mode of

PtdIns(3,5)P2 breakdown, consisting of dephosphorylation of the D3 position, leading to

PtdIns5P formation (Berger et al., 2003).

Functional data, based on PIKfyve, strongly suggests that PtdIns(3,5)P2 mainly localizes and

exerts its roles in lysosomes (Ikonomov et al., 2002; Jefferies et al., 2008; de Lartigue et al.,

2009). Indeed, inhibition of synthesis of PtdIns(3,5)P2 results in dramatic enlargement of

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lysosome-related organelles. Functionally, lack of PtdIns(3,5)P2 impairs late

endosomal/lysosomal membrane recycling and results in apparent accumulation of

autolysosomes (Rutherford et al., 2006; Ferguson et al., 2010).

The precise spatial and temporal dynamics of PtdIns(3,5)P2 during phagocytosis has remained

obscure, mostly because of a paucity of methods to detect the phosphoinositide in living cells.

Analogous to the functional effects observed in resting cells, the use of antagonists of PIKfyve

significantly impairs the transition from the early phagosome to the late stages of maturation.

This is based on a delay of the removal of PtdIns3P and the reduced accumulation of

phagolysosomal markers. Moreover, the degradative capacity of phagosomes was impaired

upon inhibition of the lipid kinase (Dayam et al., 2017).

Phosphatidylinositol 4-phosphate

PtdIns4P is the primary phosphoinositide in the Golgi apparatus. Additionally, significant levels

of the phosphoinositide are found in the cytosolic leaflet of the PM. While these two pools are

the major reservoirs of PtdIns4P in mammalian cells, functional data suggests that PtdIns4P

synthesis spans several organelles. Indeed, the development of a highly sensitive and

exquisitely specific PtdIns4P-sensing probe uncovered novel pools in the endolysosomal

system (Hammond et al., 2014).

In mammalian cells, the main PtdIns4P-synthesizing enzymes are phosphatidylinositol-4

kinases (PI4K). The PI4K family is comprised of four members—two class II members and two

class III members—each of which is responsible for generating a different pool of the lipid

(Balla, 2013; Boura and Nencka, 2015). The PM pool is generated by PI4KA (PI4KIIIα) (Balla et

al., 2008; Nakatsu et al., 2012), a cytosolic kinase that is recruited to the PM in a complex with

TTC7 and Fam126 (Chung et al., 2015a; Baskin et al., 2016; Lees et al., 2017). PtdIns4P in the

Golgi is mainly synthesized by two kinases, PI4KB (PI4KIIIβ)—in the cis Golgi—and PI4K2A

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(PI4KIIα)—in the trans Golgi—(Godi et al., 1999; Minogue et al., 2010). The latter enzyme,

additionally functions in the endolysosomal network along with its homologue PI4K2B (PI4KIIβ)

(Balla et al., 2002; Salazar et al., 2005; Jović et al., 2012).

Conversely, PtdIns4P is dephosphorylated by phosphatases containing SAC domains. In yeast

and mammalian cells, the principal PtdIns4P degrading enzyme is the highly conserved ER-

resident SAC1 enzyme (Whitters et al., 1993; Guo et al., 1999; Foti et al., 2001). Recent

studies characterized SAC2 as a PtdIns4P phosphatase in early compartments (i.e. Rab5-

positive organelles) (Hsu et al., 2015; Nakatsu et al., 2015). As described earlier,

plasmalemmal PtdIns4P can be converted into PtdIns(4,5)P2 by the action of PIP5K isoforms.

This reaction has also been proposed to occur during autophagy. Additionally, class II PI3Ks

can phosphorylate PtdIns4P at the D3 position yielding PtdIns(3,4)P2 (Misawa et al., 1998).

Finally, some evidence suggests that PLC enzymes can use PtdIns4P as a substrate for

hydrolysis (Zhang et al., 2013; Sicart et al., 2015), however their relative contribution to the

regulation of PtdIns4P levels in cells is unclear.

The longstanding notion that PtdIns4P is merely a precursor for the sequential synthesis of

plasmalemmal PtdIns(4,5)P2 and PtdIns(3,4,5)P3 has been widely disproven over the last

couple of decades (reviewed in Tan and Brill, 2014), PtdIns4P has been shown to be an

essential functional component of Golgi membranes. Indeed, PtdIns4P effectors at the Golgi

level are important for maintenance of organelle structure, vesicle budding and non-vesicular

transport. Yet, roles of the phosphoinositide in the PM and the endolysosomal system—

organelles that are central during phagocytosis—remain undefined as do PtdIns4P effectors in

these compartments.

Recent reports have suggested the presence of PtdIns4P in purified phagosomes from cell-free

systems (Jeschke et al., 2015). These PtdIns4P-positive phagosomes coincide with late

maturation stages and the presence of the lipid is dependent on PI4K2A. Moreover, blocking

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the kinase with a monoclonal antibody impairs phagolysosomal biogenesis in these cell-free

systems (Jeschke et al., 2015), resembling previous observations in autophagosomes (Wang et

al., 2015a). However, the dynamics, metabolism and functions of PtdIns4P during phagocytosis

in live cells remained unexplored when I initiated my studies.

1.6.3 Phosphoinositides during phagosome resolution

As stressed earlier, phagosome resolution is by far the least understood stage of phagocytosis.

The mechanisms responsible for it are virtually unexplored; hence it is unsurprising that

phosphoinositides with potential roles during this stage remain undefined. However, based on

the paramount functions that these lipids play in essential events during phagosome formation

and maturation, it seemed conceivable that they are also implicated in resolution.

As mentioned before, PtdIns4P is present at significant levels in the endolysosomal system

(Hammond et al., 2014). Its potential role during the final stages of phagocytosis is supported

by evidence that it may be indispensible for autolysosome and phagolysosome biogenesis

(Jeschke et al., 2015). More notably, ‘very late” fission events in resolving phagolysosomes are

characterized by vesicular and tubular structures (Krajcovic et al., 2013). These events are

likely coupled to the presentation of antigen in the cell surface to lymphoid cells. Previous

studies point to a potential role for PtdIns4P. First, in dendritic cells the adaptor protein complex

3 (AP-3) has been implicated in antigen transport to the cell surface for presentation

(Mantegazza et al., 2012). Intriguingly, there is interdependency on the localization of AP-3 and

PI4K2A to late endosomes and lysosomes (Salazar et al., 2005; Craige et al., 2008). PI4K2A is

delivered to such compartments as cargo of AP-3-generated vesicles; interestingly changes in

levels of the kinase mistarget the adaptor complex (Craige et al., 2008).

PtdIns(3,5)P2 is another potentially relevant species during phagosome resolution. The

generation of large vacuoles—originated from lysosomal compartments—is the universal

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outcome of PIKfyve inhibition (Ikonomov et al., 2002; McCartney et al., 2014). Such a

phenotype would be expected from defects in phagosome resolution due to failure to generate

post-lysosomal compartments. However, the mechanisms behind this phenomenology remain

obscure. In this context, recent evidence suggests that PtdIns5P is implicated in the ‘resolution’

of autophagosomes (Lundquist et al., 2018). This low abundance phosphoinositide is thought to

be generated—at least in part—from PtdIns(3,5)P2 by MTM phosphatases. Data suggests that

PtdIns5P then serves a precursor for the generation of PtdIns(4,5)P2 (Lundquist et al., 2018), a

species that has been implicated in autolysosome reformation (Rong et al., 2012; Du et al.,

2016). Indeed, deletion of PtdIns5P-4-kinases—enzymes responsible for the conversion of

PtdIns5P to PtdIns(4,5)P2—results in a defect in clearance of autophagosomes (Lundquist et

al., 2018). Yet, as is the case with PIKfyve inhibition, the molecular mechanisms have not been

defined.

1.7 Rationale

Mechanistically, phagocytosis is broadly divided into three major stages: phagosome formation,

maturation and resolution. While the molecular mechanisms that govern the recognition and

engulfment of particles have been extensively studied and are widely understood, comparatively

little is known about the maturation and resolution stages. In fact, only the early maturation

stage has been studied in detail. Hence much more is known about the early steps than the

concluding late stages. Moreover, as stressed earlier, the resolution stage remains virtually

unexplored. Over the last couple of decades increasing evidence has established

phosphoinositides as master regulators of phagocytosis. Indeed, phagosome formation and

early maturation are dependent on strictly coordinated localized changes in phosphoinositide

levels. These specialized phosphoinositide dynamics are responsible for the recruitment of

several effector proteins that are fundamental for phagocytosis. Hence, studying the dynamics,

metabolism and physiological implications of phosphoinositides during late phagosomal

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maturation and phagosome resolution will be of critical importance to elucidate the molecular

mechanisms behind these stages.

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Chapter 2 Hypothesis and Aims

Thesis hypothesis and aims 2

2.1 Hypothesis Due to the prominence of PtdIns(4,5)P2 and PtdIns(3,4,5)P3 in diverse signaling networks in

mammalian cells, PtdIns4P was widely regarded as a precursor molecule. The discovery of

diverse conserved roles of PtdIns4P at the Golgi apparatus established it a fundamental

signaling molecule in its own right. Importantly, the roles of PtdIns4P in the PM and in the

endolysosomal network—compartments that contain significant pools of the lipid—are yet to be

defined. Because these organelles are central during different phagocytic stages, the presence

of the phosphoinositide suggests its potential involvement in diverse aspects of the maturation

process. As described in the previous chapter, the late stages of phagocytosis are the least

understood. Specifically, the molecular mechanisms that direct the biogenesis of

phagolysosomes are poorly defined, while the resolution of phagosomes is virtually unexplored.

The paramount roles the phosphoinositides play during phagocytosis and the presence of

PtdIns4P in the endolysosomal network led me to hypothesize that fundamental events during

the late stages of phagocytosis are both regulated by and dependent on PtdIns4P.

2.2 Aims This hypothesis was examined through an experimental approach by addressing the following

aims:

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1. Determination of the dynamics of PtdIns4P during phagocytosis by live microscopy-based

techniques. This was approached through the use of genetically encoded PtdIns4P

biosensors.

2. Investigation of the PtdIns4P-metabolizing enzymes that regulate the dynamics of the

phosphoinositide during phagocytosis. In addition, I assessed their importance for the

progression of the endocytic process.

3. Definition of the potential roles of PtdIns4P during different stages of phagocytosis.

4. Elucidation of the molecular mechanisms whereby PtdIns4P exerts its functions during the

process. Specifically, I attempted to pinpoint PtdIns4P effectors at the phagosomal level.

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Chapter 3 Materials and Methods

This chapter includes material from the following:

- Fernando Montaño, Sergio Grinstein and Roni Levin. ”Quantitative phagocytosis assays

in primary and cultures macrophages.” Methods in Molecular Biology, 2018;1784:151-

163.

- Materials and Methods section of: Roni Levin, Gerald R.V. Hammond, Tamas Balla,

Pietro de Camilli, Gregory, D. Fairn and Sergio Grinstein. “Multiphasic dynamics of

phosphatidylinositol 4-phosphate during phagocytosis.” Molecular Biology of the Cell,

2017, Jan 1;28(1):128-140.

- Materials and Methods section of: Roni Levin, Tal Keren-Kaplan, Braeden Ego,

Fernando Montaño, Jess Diciccio, William S. Trimble, Michael C. Bassik, Juan

Bonifacino, Gregory D. Fairn and Sergio Grinstein. “Phagolysosome resolution is

mediated by contacts with the endoplasmic reticulum and PtdIns4P down-regulation”.

Submitted

General methods 3 3.1 Introduction

The signaling that triggers phagocytosis must be highly coordinated in space and time.

Additionally, due to its unsynchronized and transient nature, traditional biochemical methods

that depend on populations of large numbers of cells are inadequate to study detailed molecular

aspects of phagocytosis. Unlike population-based assays—which have been highly informative

in their own right—, single cell assays provide high spatial and temporal resolution (Swanson,

2004). The generation of sophisticated molecular biology tools have enabled the development

of non-invasive and continuous measurements in live cells (Lu et al., 2017). These methods,

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coupled with specialized microscopy techniques and image analysis software, have given rise to

powerful combinations to investigate phagocytosis at the molecular level.

It is worth noting that phagocytosis is an umbrella term that describes a family of processes that,

while phenotypically related, differ at the molecular level. Individual ligand-receptor pairs trigger

distinct signaling cascades that, while ultimately causing internalization, reach this goal by

different means (Flannagan et al., 2014; Lew et al., 2014). Moreover, even when the same

receptors are engaged, the size, shape and rigidity of the targets influences signaling and its

integration (Cox et al., 1999b; Champion and Mitragotri, 2006). Because of such caveats, it has

been essential to develop quantitative measurements to analyze various aspects of

phagocytosis. Here we describe methods to quantify basic and advanced features of the

phagocytic response.

Whenever possible, studying phagocytosis in primary cells is preferable; their responsiveness is

unparalleled. Reactions such as phagosome acidification and the generation of reactive oxygen

species are prominent and can be readily measured in primary phagocytes (Balce and Yates,

2017). Moreover, antibodies are available for detection of specific components by

immunofluorescence. However, when genetically-encoded tools and routine genetic

manipulations are required, the use of cell lines—such as immortalized murine macrophage-like

cells—is recommended (Flannagan and Grinstein, 2010). Using such cells bypasses the need

to isolate (and differentiate) primary phagocytes, a time-consuming and labor-intensive

procedure.

3.2 Reagents

Sheep erythrocytes (SRBC) (10% suspension) were purchased from MP Biomedicals. Anti-

sheep red blood cell antibodies were purchased from Cedarlane Laboratories. Polystyrene

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microspheres (1.6 µm in diameter) functionalized with divinylbenzene were obtained from Bangs

Laboratories. Rapamycin, human IgG, cresyl violet acetate and wortmannin were from Sigma-

Aldrich. Fluorescent antibodies against mouse and rabbit were from Jackson ImmunoResearch

Labs. Paraformaldehyde (16% w/v) was from Electron Microscopy Sciences.

3.3 Cell culture

RAW 264.7 and COS-1 cells were obtained from the American Type Culture Collection

(Manassas, VA). RAW 264.7 murine macrophages were routinely grown in T-25 tissue culture

flasks in 10 mL of Roswell Park Memorial Institute medium (RPMI) (Wisent; St. Bruno, Qc)

supplemented with 10% heat inactivated fetal bovine serum (HI-FBS) (Wisent) at 37°C under

5% CO2. Upon reaching 70-80% confluence, cells were washed with pre-warmed sterile

phosphate-buffered saline (PBS) (Wisent; St. Bruno, Qc). After removal of the PBS, fresh RPMI

plus 10% HI-FBS was added. Cells were then lifted by gently scraping, transferred into a 15 ml

conical tub and spun down at 300 x g for 3 min. Next that supernatant was removed and the

pellet resuspended in pre-warmed RPMI plus 10% HI-FBS. To passage the culture, cells were

added to a new T-25 tissue culture flask containing pre-warmed RPMI plus 10% HI-FBS in a 1:5

dilution. Alternatively, resuspended cells were seeded on glass coverslips contained in 12-well

tissue culture plates with pre-warmed RPMI plus HI-FBS. Cells were then incubated at 37°C

under 5% CO2.

Generation of COS-1 cells stably expressing FcγRIIa (COS-1-FcγRIIa) were previously

generated and described (Downey) COS-1-FcγRII were routinely grown in Dulbecco’s Modified

Eagle’s Medium (DMEM) (Wisent) supplemented with 10% HI-FBS. Cell passage and seeding

were performed as for RAW 264.7 cells, however COS-1 cells were lifted using trypsin (Wisent;

St. Bruno, Qc) as opposed to scraping.

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3.4 Primary cell isolation and differentiation

Primary human monocytes were isolated from peripheral blood of healthy donors. In brief,

blood was diluted with PBS (1:1) and overlaid on a Lympholyte-H separation density gradient

(Cedarlane labs; Burlington, ON) in 50 ml conical tubes. The tubes were then spun down at 800

x g for 20 min. without acceleration and deceleration. Then, the layer containing the monocytes

/ macrophages was extracted from the gradient and resusupended in Hank’s Balanced Salt

Solution (HBSS) (Wisent). Cells were then spun down at 500 x g for 10 min. and then

resuspended in RPMI + 10% HI-FBS + penicillin and streptomycin (P/S). The resuspended

cells were transferred into a 10 cm dish and incubated 37°C under 5% CO2.

For macrophage differentiation, after 1 h of incubation, media was replaced with RPMI + 10%

HI-FBS + P/S + macrophage colony stimulating factor (MCSF) (25 ng/ml). This media was

replaced and replenished every two days. Six days after isolation and treatment with MCSF,

the adherent cells were fully differentiated macrophages. To seed macrophages onto glass

coverslips, cells were washed with PBS and then lifted with acutase (Invitrogen; Carlsbad, CA)

for 20 min. Cells were then added to coverslips within wells containing pre-warmed media.

3.5 Antibodies

Mouse monoclonal anti-PI4K2A was a kind gift from T. Balla (National Institutes of Health;

Bethedsa, MD); Rabbit monoclonal anti-ORP1L for immunofluorescence was a kind gift from V.

Olkkonen; rabbit polyclonal anti-ORP1L for western blots (1:2000, Abcam); goat polyclonal anti-

VAPA (1:200, Santa Cruz,); rabbit polyclonal anti-VAPB (1:500, Sigma); mouse monoclonal

anti-GAPDH (1:5000, Millipore,); anti-GFP (1:500, Santa Cruz).

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3.6 Plasmids

The following plasmids used in this study have been previously described: GFP-2xP4M-SidM,

GFP-P4M-SidM, iRFP-FRB-Rab7, GFP-PI4KB, GFP-PI4K (Hammond et al., 2014); GFP-

PI4K2A, GFP-PI4K2B (Balla et al., 2002); mRFP-PH-PLC∂ (Stauffer et al., 1998); mRFP-

2xFYVE-EEA1, GFP-PH-Gab2 (Bohdanowicz et al., 2010), mRFP-Rab5, mRFP-Rab7 (Vieira et

al., 2003); GFP-Sac2 (Nakatsu et al., 2015); mRFP-FKBP-Sac1 (Hammond et al., 2012);

mRFP-FKBP (Szentpetery et al., 2010); GFP-RILP-C33 (Cantalupo et al., 2001); Lyn11-FRB

(Bohdanowicz et al., 2013); mCh-sialyltransferase was a kind gift from Dr. E. Rodriguez

Boulan; GST-GFP-P4M was generated using GFP-P4M as a template and the following

primers: forward: 5-gcggaattcatgtctaaaggtgaagaatt-3’; reverse: 5’gcgctcgagttattttatcttaatggtttg-

3’. The PCR product was digested with EcoRI and Xhol and cloned into a pGEX-6P1; mRFP-

FKBP-PLCβ3 was generated using the constitutively active DXY-PLCb3 template kindly

provided by T.K. Harden and J. Sondek, (University of North Carolina School of Medicine,

Chapel Hill, North Carolina) (Charpentier et al., 2014) and using primers: forward: 5’-

atatcgatcggtccaggcgctgcagttgg-3’; reverse: 5’-atattctagatcaaatgtagtccgaggcttcggtgtagatg-3’ for

PCR amplification. The PCR product was digested with Pvu and XbaI and cloned into the

mRFP-FKBP12-5-ptase domain construct obtained from plasmids amplified in Dam- bacterial

strains and digested with the same restriction enzymes to replace the 5-ptase domain with the

PLCb3 construct; GFP in GFP-2xP4M was digested using NheI and BglIII, and was replaced

with mCherry to generate mCh-2xP4M-SidM. The following plasmids were kind gifts: GFP-

ORP1L and GFP-ORP1L-D478A from J. Neefjes; mCh-ORP1L from N. Ridgway; CFP-Rab7

from J. Brumell; GFP-VapA, GFP-VAPB, mCh-VapA and mCh-VAPB from W. Trimble, GFP-

PH-PLC∂ from T. Meyer. The following plasmids were previously described: mCh-Arl8B; RFP-

Rab7-T22N; the following plasmids were generated using InFusion® HD EcoDry™ (Clontech,

Mountain View, CA): mCh-ORP1L-HH651/652AA and GFP-ORP1L-HH651/652AA using the

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following primers: CTCCGAACAGGTCAGCGCTGCCCCACCAATCAGTGCAT and

ATGCACTGATTGGTGGGGCAGCGCTGACCTGTTCGGAG.

3.7 Transient transfections

For transient transfections, ≈80% confluent monolayers of RAW 264.7 cells were lifted by

scraping and plated onto 1.8 cm glass coverslips at a density of 5 × 104 cells per coverslip.

Macrophages were allowed to recover for 18–24 h and then transfected with FuGENE HD

(Promega) according to the manufacturer's instructions. Briefly, 1 µg of plasmid DNA and 3 µL

of the transfection reagent were mixed in 100 µL of serum-free RPMI and incubated for 15 min.

The mix was then distributed equally into four wells of a 12-well plate containing the cells. Cells

were typically used for experiments 12-18 h after transfection.

3.8 Particle opsonization

SRBC were centrifuged from a 10% suspension and washed 2X with PBS. Then, cells were

resuspended in 200 µl of PBS and 4 µl of rabbit IgG fraction against SRBC was added. Next,

the mix was incubated in a heated shaker at 1050 RPM and 37°C for 1 h. SRBC were then

washed 2X with PBS and finally resuspended in 1 ml of PBS. For SRBC labeling, a

fluorescently conjugated secondary antibody against rabbit was added (1:1000) and the cells

were incubated under gentle rotation at RT for 40 min.

For human RBC (HRBC), the procedure was repeated with specific differences. HRBC were

obtained from peripheral blood of healthy donors. Cells were opsonized with mouse IgG

fraction against HRBC. Finally, cells were labeled with a fluorescently conjugated secondary

antibody against mouse.

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3.9 Phagocytosis assay

For all phagocytosis assays, ≈ 5 × 104 cells were seeded onto 1.8 cm glass coverslips and allowed to

grow for 18-24 hrs. Phagocytosis was initiated by adding 15 µL of a 10-fold dilution of the IgG-SRBC or

IgG-bead suspension to individual wells of a 12-well plate or directly into an imaging chamber containing

a coverslip with cells. In most cases phagocytosis was synchronized by sedimenting the particles by

centrifugation (300 x g for 10 sec).

3.10 Gene silencing

The siRNAs (27mer -dicer substrate) targeting mouse Sac2/Innp5F were from Origene. The

targeting sequences were siRNA1: 5'-GGAAUGCGGUAUAAACGAAGAGGAG-3' and siRNA2:

5'-ACAAGCCUGAGAAGAUUAUACCATC-3'. To silence PI4K2A, siRNA targeting mouse

Pi4k2a was purchased from Dharmacon. The targeting sequences: 5'-

UGAGGGAGCCUGUUAUCAA-3', 5'-GGACACAGAUUGGGUGAUG-3', 5'-

GAAUCGGCCUGCCACCAAA-3', 5'-GAGACGAGCCCGCUAGUGU-3', 5'-

CUACAAAGAUGCAGACUAUUU-3' and 5'-CCAGAUGCCACCUGUGAUUUU-3'.

The siRNA1 (5'-UGAAGCAGAACCUCUUCCUGAUU-3') previously used by (Wang, Balla,

Jovic) and the siRNA2 (5'-CUACAAAGAUGCAGACUAUUU-3') targeting PI4K2A in COS-1 cells

were also from Dharmacon. Lastly, the siRNA oligos (Stealth RNAi™) against mouse Orp1l

(Osbpl1a) were from ThermoFisher Scientific (Waltham, MA) with the following targeting

sequences: GGCCAUGGACUUGAAGGAGUCGUUA and

GCAUCCUUAGUGAGGAGGAGUUCUA. Oligonucleotides were delivered into RAW

macrophages by electroporating 5 × 105 RAW 264.7 cells with 200 pmol of the siRNA pool with

the Neon transduction system (ThermoFisher Scientific), using a single 20-millisec pulse of

1750 V. Electroporated cells were allowed to recover for 24 h before being lifted once again for

a second round of electroporation. Knockdown efficiency and phagocytosis were assessed 48

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h after the initial electroporation. When transfection of plasmid DNA was required, cells were

re-plated in 12-well plates after the second round of silencing and allowed to recover in fresh

medium for 8 hrs. Then, cells were transfected as above and used for experiments after an

additional 18-24 h incubation.

Alternatively, Oligonucleotides were delivered to COS-1 cells with HiPerFect transfection

reagent (Qiagen) following the manufacturer’s instructions. Cells were plated in six-well plates

24 h before silencing. Then, cells were treated with a mix of the reagent and oligonucleotides

for 18 h and allowed to recover in fresh medium for 6 hrs. A second round of silencing was

performed for 18 hrs. Cells were allowed to recover for 30 h before experiments.

When cells had to be transfected, they were re-plated in 12-well plates after the second round of

silencing and allowed to recover in fresh medium for 8 hrs. Then, cells were transfected as

above and used for experiments 18-24 h after transfection.

3.11 Quantitative RT-PCR

Total RNA was extracted and isolated using the GeneJET RNA Purification kit (ThermoFisher

Scientific). For cDNA generation, equivalent amounts of total RNA were loaded to use the

SuperScript VILO cDNA kit (ThermoFisher Scientific). Then, qPCR reactions were performed in

96-well plates using TaqMan reagents (ThermoFisher Scientific) on a Step One Plus Real-Time

PCR thermal cycler with Step One software (v2.2.2; Applied Biosystems). Assays for reference

gene and target gene were duplexed in triplicate for every experimental replicate. The TaqMan

assays were as follows. In RAW macrophages: Abt1 (reference gene) Mm00803824_m1; and

Osbpl1a (Orp1l) Mm00498552_m1; Pi4k2A: Mm01197215_m1; and Inpp5f (Sac2:

Mm00724391_m1). In COS-1 cells: reference gene: CDKN1A: Hs00355782_m1; and target

gene PI4K2A: Hs00218300_m1.

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3.12 Generation of CRISPR KO cell lines

ORP1L KO CRISPR cell lines were generated as follows: RAW 264.7 macrophages stably

expressing EF1Alpha-Cas9-BFP were lentivirally transduced with constructs expressing a

puromycin resistance cassette and an ORP1L-targeting sgRNA. 72 hours following infection,

selection was performed with puromycin (10 ug/mL) for 5 days. Following selection, single cells

were sorted into 96-well plates and expanded for two to three weeks to generate single cell

clonal knockout lines.

3.13 Gene editing measurements by Sanger sequencing

Total genomic DNA was isolated from cells using QuickExtract DNA Extraction Solution (VWR,

Radnor, PA, cat# QE09050). For the PCR reaction, primers were designed approximately 250-

300 basepairs upstream and 450-550 basepairs downstream of the predicted cut site.

Following PCR reaction completion using a C1000 Touch Thermo Cycler (Bio-Rad), PCR

products were run on an agarose gel and were gel-purified over an Econospin DNA column

(Epoch, Missouri City, TX, cat# 1910-250) using a QIAquick Gel Extraction Kit (Qiagen, Hilden,

Germany, cat# 28706). Sanger sequencing ab1 data were obtained from Quintara Biosciences

(San Francisco facility) and the editing efficiency and indels of knockout cell lines were

assessed using the online TIDE analysis tool (https://tide.deskgen.com/).

3.14 Detection of acidification and lysosomal labeling

Cells were treated with cresyl violet acetate (1 µM final concentration) for 2 min, then washed

twice with PBS and imaged. Lysosomes were loaded with 0.1 mg/mL dextran for 3 h at 37°C

(5% CO2 balance air), followed by a chase period of at least 30 min.

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3.15 Confocal microscopy

For imaging, cells grown on glass coverslips, were mounted in a Chamlide magnetic chamber

(Seoul, Korea), overlaid with 1 mL of HBSS (Wisent Bioproducts) and maintained at 37ºC.

Fluorescence microscopy was performed using spinning-disk confocal microscopes (Quorum

Technologies). Our systems are based on an Axiovert 200M microscope (Carl Zeiss) equipped

with a ×63 oil-immersion objective (NA 1.4) and a x 1.5 magnifying lens. The microscopes carry

a motorized XY stage (Applied Scientific Instrumentation), a Piezo Z-focus drive and diode-

pumped solid-state lasers emitting at 440, 491, 561, 638 and 655 nm (Spectral Applied

Research). Images were recorded with back-thinned, cooled charge-coupled device cameras

(Hamamatsu Photonics) under command of the Volocity software (version 6.2.1; PerkinElmer).

3.16 Lattice light-sheet microscopy

The lattice light-sheet microscope (LLSM) used in these experiments is housed in the Advanced

Imaged Center (AIC) at the Howard Hughes Medical Institute Janelia research campus with the

support of John Heddleston, Satya Khuon and Teng Leong-Chew. The system is configured

and operated as previously described (Chen et al., 2014). Briefly, RAW macrophages were

grown on 5 mm round glass coverslips (Warner Instruments, Catalog # CS-5R) ~36 h before

experiments. After adhesion, cells were transfected with Fugene HD 12 – 18 h before imaging

as described above. For the assay, IgG-RBC were added to the wells containing cells on

coverslips. The plates were spun down (300Xg for 10 s) in order to synchronize phagocytosis.

Immediately after, coverslips were placed on a custom-made stainless-steel holder. Imaging

was conducted in HBSS at 37 °C. Samples were illuminated by a 2D optical lattice generated

by a spatial light modulator (SLM, Fourth Dimension Displays). The sample was excited by 488

nm or 560 nm diode lasers (MPB Communications) at 25% AOTF transmittance and 50 mW

initial box power through an excitation objective (Special Optics, 0.65 NA, 3.74-mm WD).

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Fluorescent emission was collected by detection objective (Nikon, CFI Apo LWD 25XW, 1.1

NA), and detected by a sCMOS camera (Hamamatsu Orca Flash 4.0 v2). Acquired data were

deskewed as previously described (Chen et al., 2014) and deconvolved using an iterative

Richardson-Lucy algorithm. Point-spread functions for deconvolution were experimentally

measured using 200nm tetraspeck beads adhered to 5 mm glass coverslips (Invitrogen, Catalog

# T7280) for each excitation wavelength.

3.17 Transmission electron microscopy

RAW macrophages were cultured on 18-mm coverslips and challenged with IgG-SRBC as

described before. Cells were incubated following phagosome maturation until pre-determined

time points, fixed with 2% (vol/vol) glutaraldehyde and processed for TEM using standard

methods.

3.18 Image processing

For confocal microscopy, images were acquired using Volocity (Perkin Elmin, Woodbridge, ON)

and exported to ImageJ (National Institutes of Health, Bethesda, MD) for analysis, quantification

and contrast enhancement. Selection of regions of interest, fluorescence intensity

measurements and brightness/contrast corrections were performed with ImageJ. Brightness

and contrast parameters were adjusted across entire images and without altering the linearity of

mapped pixel values. For LLSM 3D data visualization and analysis images were exported to

Imaris (Bitplane, Concord, MA).

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3.19 Protein purification

BL21 competent cells were transformed with GST tagged proteins. A single transformed colony

was inoculated in 100 ml of Terrific Broth (Invitrogen) supplemented with 0.1% glycerol and

incubated overnight at 37°C. The next day, 20 ml of the overnight culture were inoculated in

400 ml of Terrific Broth with glycerol and incubated for 2 h at 37°C. After the incubation,

isopropyl β-D-1-thiogalactopyranoside (Sigma) was added (final concentration 0.25mM) and the

culture was incubated at 30°C for 4 h. After the incubation, cells were lysed using FastBreakTM

cell lysis reagent (Promega) according to manufacturer’s instructions. The cell lysate was then

centrifuged and the supernatant was collected. The presence of GST-GFP-P4M was confirmed

through western blot using an α-GFP antibody [GFP(B-2):sc-9996, Santa Cruz Biotechnology]

detecting a band with a size ≈65 kDa, corresponding to the added molecular weight of GST,

GFP and P4M. The supernatant was then incubated with Glutathion SepharoseTM 4B (GE

Healthcare) (previously washed with binding buffer [PBS, pH 7.3] three times), overnight at 4°C.

The solution was then centrifuged and washed three times with binding buffer. Lastly, the pellet

was incubated in elution buffer (50 mM Tris-HCL 10 mM reduced glutathione, pH 8.0) for 30 min

at RT and the elution was collected. The presence of the purified GST-GFP-P4M was

confirmed through western blot using an α-GFP antibody [GFP(B-2):sc-9996, Santa Cruz

Biotechnology].

3.20 Protein-lipid overlay assay

A PIP strip (Echelon Biosciences) was blocked overnight at 4°C with PBS + 0.1% tween (PBS-

T) + 3% BSA, with agitation. The next day, the membrane was covered with purified GST

tagged protein (150 pmol/ml in blocking solution), and incubated overnight at 4°C with agitation.

The next day, the membrane was washed with PBS-T for 20 min with agitation five times. After

the washes, the PIP strip was covered with either α-GFP or α-GST antibody diluted in blocking

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buffer (1:500) and incubated for 1 h with agitation at RT. After the incubation, the membrane

the strip was washed as described above. Next, the strip was incubated in a solution containing

a α-mouse HRP-conjugated antibody (Jackson ImmunoResearch Labs) for 1 h with agitation at

RT. The membrane was then washed as described above. Lastly, the detection step was

performed using the AmershamTM ECLTM Prime Western Blotting Detection Reagent (GE

Healthcare) according to manufacturer’s instructions.

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Chapter 4 Multiphasic Dynamics of Phosphatidylinositol 4-phosphate During

Phagocytosis This chapter has been modified from the following: Roni Levin, Gerald R.V. Hammond, Tamas

Balla, Pietro de Camilli, Gregory, D. Fairn and Sergio Grinstein. “Multiphasic dynamics of

phosphatidylinositol 4-phosphate during phagocytosis.” Molecular Biology of the Cell, 2017, Jan

1:28(1):128-140.

Multiphasic dynamics of phosphatidylinositol 4-4phosphate during phagocytosis

4.1 Abstract

We analyzed the distribution, fate and functional role of phosphatidylinositol 4-phosphate

(PtdIns4P) during phagosome formation and maturation. To this end we used genetically-

encoded probes consisting of the PtdIns4P-binding domain of the bacterial effector SidM.

PtdIns4P was found to undergo complex, multiphasic changes during phagocytosis. The

phosphoinositide, which is present in the plasmalemma prior to engagement of the target

particle, is transiently enriched in the phagosomal cup. Shortly after the phagosome seals,

PtdIns4P levels drop precipitously due to the hydrolytic activity of Sac2 and phospholipase C,

becoming undetectable for ≈10 min. PtdIns4P disappearance coincides with the emergence of

phagosomal PtdIns3P. Conversely, the disappearance of PtdIns3P that signals the transition

from early to late phagosomes is accompanied by resurgence of PtdIns4P, which is associated

with the recruitment of phosphatidylinositol 4-kinase 2A. The reacquisition of PtdIns4P can be

prevented by silencing expression of the kinase and can be counteracted by recruitment of a 4-

phosphatase with a heterodimerization system. Using these approaches we found that the

secondary accumulation of PtdIns4P is required for proper phagosomal acidification. Defective

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acidification may be caused by impaired recruitment of Rab7 effectors, including RILP that was

shown earlier to displace phagosomes towards perinuclear lysosomes. Our results show

multimodal dynamics of PtdIns4P during phagocytosis and suggest that the phosphoinositide

plays important roles during the maturation of the phagosome.

4.2 Introduction

Phagocytosis, the ingestion of particulate matter, is essential for the elimination of invading

pathogens, serving as a first line of defense of the immune system (Levin et al., 2016). It also

plays a key role in tissue homeostasis and remodeling, disposing of apoptotic bodies and debris

(Elliott and Ravichandran, 2010). The efficient and timely removal of effete cells is necessary to

prevent secondary necrosis and unwanted inflammation.

Phagocytosis is initiated by the engagement of surface receptors that trigger extensive

remodeling of the plasma membrane (PM) and the actin cytoskeleton (Levin et al., 2016),

culminating with the extension of pseudopods that surround and engulf the target. The nascent

phagosome then undergoes profound changes, acquiring microbicidal and degradative

properties that enable it to process its contents. This is accomplished through a series of fission

and fusion events with other endomembrane compartments, a process globally termed

phagosome maturation (Fairn and Grinstein, 2012).

Phagosome formation and maturation are both temporally and spatially restricted; as such, they

are subject to exquisite control, relying on highly coordinated signals. Phosphatidylinositol

derivatives are central to the signaling sequence (Swanson, 2014; Levin et al., 2015). A

combination of biochemical determinations and imaging using genetically-encoded biosensors

has revealed that phosphatidylinositol 3,4,5-trisphosphate [PtdIns(3,4,5)P3] accumulates at the

phagosomal cup, while phosphatidylinositol 4,5-bisphosphate [PtdIns(4,5)P2] is acutely depleted

(Botelho et al., 2000; Marshall et al., 2001). These changes orchestrate actin remodeling.

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Shortly after sealing, the phagosome accumulates phosphatidylinositol 3-phosphate (PtdIns3P),

which signals the initiation of the maturation process (Vieira et al., 2001).

Unlike these phosphoinositides, little is known about phosphatidylinositol 4-phosphate

(PtdIns4P) in phagocytosis, despite its abundance. This is due, at least partly, to the lack of

appropriate tools to detect this lipid with sufficient sensitivity and specificity. This hurdle,

however, was recently overcome by the development of a reliable biosensor to monitor

PtdIns4P in live cells. Hammond et al. (Hammond et al., 2014) developed a novel PtdIns4P-

sensing probe utilizing a domain of the Legionella pneumophila PtdIns4P-interacting effector

SidM (Brombacher et al., 2009) that recognizes the headgroup of the phospholipid

stereospecifically. Expression of fluorescently-tagged versions of this biosensor in mammalian

cells not only highlighted the previously identified pools of PtdIns4P in the Golgi and plasma

membrane, but also revealed the presence of PtdIns4P in Rab7-positive late endosomes and

lysosomes (Hammond et al., 2014).

We took advantage of this newly developed probe to analyze the fate of PtdIns4P during

phagosome formation and maturation. Our results revealed the occurrence of striking triphasic

changes in the content of PtdIns4P of the nascent and maturing phagosome. Importantly, we

detected a previously unappreciated wave of PtdIns4P regeneration that is critical for the

transition from early- to late-phagosomes, enabling luminal acidification and the acquisition of

the Rab7-interacting lysosomal protein (RILP).

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4.3 Results

4.3.1 Detection of PtdIns4P in macrophages

We initially assessed the ability of a chimeric construct consisting of the P4M domain of SidM

linked to GFP (GFP-P4M) to detect PtdIns4P in macrophages. When transiently expressed in

RAW 264.7 cells—a murine line of monocyte/macrophage origin—GFP-P4M was most

prominently localized to the Golgi apparatus / trans-Golgi network (TGN), with less prominent

accumulation at the PM and in cytoplasmic puncta (Fig. 4.1ai). However, a large fraction of the

probe was unbound (cytosolic), lowering the contrast and making the organellar pools difficult to

discriminate and track during phagocytosis. We therefore tested a second probe consisting of

two P4M domains fused in tandem and tagged with GFP (GFP-2xP4M), which is expected to

bind to PtdIns4P-containing membranes with greater avidity. GFP-2xP4M labeled not only the

Golgi elements but also the PM (Fig 4.1aii) and endosomal structures that were Rab7-positive

(not shown). We observed far less cytosolic GFP signal in cells expressing the tandem probe

compared to the single domain, consistent with the notion that GFP-2xP4M binds PtdIns4P with

higher avidity. Of note, the morphology of the Golgi apparatus, assessed using mCh-

sialyltransferase as marker, was not affected in cells expressing low to medium levels of the

tandem biosensor (Fig. 4.2) as opposed to the distorted Golgi observed with high expression

(Hammond et al., 2014). Additionally, we observed no perturbations in the distribution of a

PtdIns(4,5)P2-binding probe (PH-PLCδ), suggesting that the metabolism of this lipid was not

altered. We therefore concluded that, when expressed at moderate levels, the GFP-2xP4M

probe did not interfere noticeably with overall phosphoinositide metabolism or cell function.

The specificity of 2xP4M for PtdIns4P, and hence the validity of the determinations, was tested

next. To this end we constructed a GST-GFP-P4M plasmid to generate and purify recombinant

protein. We used hydrophobic membranes spotted with different lipid species to test the

interaction of the purified protein with biologically relevant lipids. As shown in Fig. 4.1b,

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Figure 4.1. PtdIns4P undergoes triphasic changes during phagocytosis. a) Confocal section of RAW 264.7 cells transiently expressing (i) GFP-P4M or (ii) two P4M domains fused in tandem tagged with GFP (GFP-2xP4M). b) Protein-lipid overlay assay of purified GST-GFP-P4M and 100 pmol of the indicated lipids spotted in a hydrophobic membrane. The protein was detected with an α-GFP antibody followed by an HRP-conjugated secondary antibody. c) Schematic diagram illustrating the recruitment of Sac1S.c to the PM through a rapamycin heterodimerization system. d) RAW 264.7 cells transiently co-

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expressing GFP-2xP4M, mCh-FKBP-Sac1 and Lyn11-FRB were imaged before (left) and after (right) the addition of rapamycin (1 µM); insets show mCh-FKBP-Sac1 (F-Sac1) fluorescence (black and white were inverted for clarity). e) Time course of the changes in the ratio between the plasmalemmal and the Golgi GFP-2xP4M intensities upon the recruitment of the 4-phosphatase Sac1 to the PM. Time “0” represents values before the addition of rapamycin. Values are means ± standard error of the mean (SEM) of four independent experiments. f) Time-lapse gallery of confocal micrographs acquired during phagocytosis of TAMRA-labeled IgG-opsonized sheep red blood cells (IgG-SRBCs; shown in red) by RAW 264.7 cells transiently expressing GFP-2xP4M (green). The panels indicate the time elapsed from the moment the target was engaged. Insets show magnifications of the area delimited by dotted white boxes. g) Summary of the changes in PtdIns4P content of phagosomes following engagement of IgG-SRBCs. The intensity of GFP-2xP4M in phagosomes was quantified and normalized to that of the plasmalemma. Three phases were defined: I) an initial transient increase (0 - 2 min); II) a subsequent disappearance (2 - 10 min); and III) a reappearance and gradual increase (10 - 30 min). Data are expressed relative to the maximum value attained during phase I. Values are means ± SEM of 25 phagosomes from 15 independent experiments. h) PtdIns4P reappearance in phagosomes was often accompanied by formation of 2xP4M-positive tubules. Scale bars = 5 µm.

GST-GFP-P4M bound PtdIns4P with exquisite specificity. This finding is consistent with the

observations made using full-length SidM (Brombacher et al., 2009), implying that selectivity for

PtdIns4P was retained following excision of the P4M domain and attachment to GFP. Next, we

assessed the specificity of the sensing probe in cells. To verify the specificity of the probe when

expressed in macrophages we depleted PtdIns4P from the PM in a controlled manner, using a

heterodimerization system to recruit the PtdIns4P-specific phosphatase SAC1 using rapamycin

(Fig. 4.1c). Cells were triple-transfected with the chimera mCh-FKBP-SAC1, a construct

containing FRB targeted to the PM through the N-terminal 11 amino acids of Lyn kinase (Lyn11-

FRB), and with GFP-2xP4M. As illustrated in Fig. 4.1d, a significant fraction of GFP-2xP4M

localized to the PM before the addition of rapamycin, while mCh-FKBP-SAC1 remained soluble

in the cytosol (Fig. 4.1d left). Upon addition of rapamycin mCh-FKBP-SAC1 rapidly translocated

to the PM, catalyzing the hydrolysis of PtdIns4P and causing the release of the GFP-2xP4M

probe within seconds; the biosensor released from the PM then accumulated on the Golgi and

endosomal membranes (Figs. 4.1d and e). Taken together these results validate 2xP4M as a

specific and reliable biosensor for PtdIns4P in macrophages.

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Figure 4.2. 2xP4M expression does not affect Golgi morphology. a) Representative confocal micrograph of a RAW 264.7 cell co-expressing the PtdIns4P-biosensor GFP-2xP4M and the Golgi-marker mCh-sialyltransferase. Main panel shows mCh-sialyltransferase fluorescence; the cell outline is drawn in yellow. Bottom left inset shows GFP-2xP4M fluorescence. b) Representative confocal micrograph of a RAW cell expressing only the Golgi-marker mCh-sialyltransferase. Top right insets show magnifications of the corresponding dotted white boxes. Scale bars = 5 µm.

4.3.2 PtdIns4P dynamics during phagosome formation and maturation

We proceeded to determine the dynamics of PtdIns4P during phagocytosis. We chose FcγR-

mediated phagocytosis, the most extensively characterized phagocytic system, as an

experimental model. IgG-SRBCs were used as phagocytic targets and the dynamics of the

PtdIns4P biosensor were monitored by time-lapse video microscopy. As illustrated in Fig. 4.1f

and quantified in Fig. 4.1g, when RAW 264.7 cells transiently expressing GFP-2xP4M were

challenged with tetramethylrhodamine (TMR)-labeled IgG-SRBCs, the biosensor accumulated

in the forming phagocytic cup, relative to unengaged areas of the plasmalemma. Upon

phagosome closure (approximately 2 min after initiation of internalization) the biosensor levels

peaked (end of phase I in Fig. 4.1e). Strikingly, seconds after this marked increase, GFP-

2xP4M detached from the sealed phagosome and remained absent from the compartment for

approximately 8-9 min. Furthermore, the biosensor reappeared gradually in phagosomes,

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starting ≈10 min after the particle initially contacted the macrophage (Figs. 4.1f and g). The

levels of GFP-2xP4M associated with the phagosome increased continuously for the next 15

min, surpassing the initial maximum observed upon closure and remaining high for at least 30

min after the initiation of phagocytosis (Figs. 4.1f and g). The reappearance of PtdIns4P was

often accompanied by the formation of dynamic 2xP4M-positive tubules and vesicles that

emanated from phagosomes towards the juxtanuclear region (Fig. 4.1h). Based on similar

observations made in over 80 phagosomes from multiple cells and preparations we defined a

triphasic pattern of PtdIns4P dynamics: I) a transient increase at the phagocytic cup during

phagosome formation, II) a virtually complete disappearance from the early sealed phagosome

and III) a gradual reappearance on maturing phagosomes that ultimately surpasses the density

at the PM (Fig. 4.1g).

The mechanisms underlying the changes in PtdIns4P were analyzed next. Conversion to other

inositide species seemed likely to account for the observed changes. We therefore compared

the dynamic changes of PtdIns4P with those of PtdIns(4,5)P2. As reported earlier (Botelho et

al., 2000), when visualized using mRFP-PH-PLCδ, PtdIns(4,5)P2 underwent a transient

accumulation shortly after the macrophages engaged the target, followed by a precipitous

decrease that was obvious even before phagosomal closure was completed (Figs. 4.3a and c).

PtdIns(4,5)P2 remains undetectable thereafter, failing to reappear throughout the maturation

period. Comparison of the changes in PtdIns4P and those of PtdIns(4,5)P2 in cells co-

transfected with GFP-2xP4M and mRFP-PH-PLCδ revealed that: a) the initial increase in

PtdIns(4,5)P2 does not incur depletion of PtdIns4P, which in fact also increases during the

earliest stages of particle engagement; b) the steep initial decrease in in PtdIns(4,5)P2 is not

accompanied by significant changes in PtdIns4P, suggesting that other mechanisms, such as

hydrolysis to diacylglycerol by phospholipase C are involved; c) the final stage of PtdIns(4,5)P2

disappearance coincides with a significant increase in PtdIns4P, consistent with hydrolysis by 5-

phosphatases and d) the sharp decrease in PtdIns4P observed after the phagosome seals

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Figure 4.3. Comparison of the changes in PtdIns4P content to those of PtdIns(4,5)P2 and PtdIns3P. a) Time-lapse gallery of confocal micrographs of RAW 264.7 cells co-expressing GFP-2xP4M and mRFP-PH-PLCδ, a PtdIns(4,5)P2 biosensor, during phagocytosis of IgG-SRBCs. b) Time-lapse gallery of confocal micrographs of cells co-expressing GFP-2xP4M and mRFP-2xFYVE-EEA1, a PtdIns3P biosensor, during phagocytosis of IgG-SRBCs. In a. and b. the insets show magnifications of the area delimited by dotted white boxes; the dotted circles in the insets show the location of the phagosome. Scale bars in a. and b. = 5 µm. c) Time course of the changes in PtdIns4P and PtdIns(4,5)P2 during phagosome formation. The intensity of GFP-2xP4M fluorescence in the phagosome was measured and normalized to the plasmalemmal intensity; to enable comparison between cells and between experiments the data are expressed relative to the maximum value (green line, white circles). A similar analysis was applied to mRFP-PH-PLCδ (red line, black circles), a PtdIns(4,5)P2 biosensor. d) Time course of the changes in PtdIns4P and PtdIns3P during phagosome formation and maturation. PtdIns4P was monitored as above, using GFP-2xP4M. mCh-FYVE-EEA1 was used as a PtdIns3P probe; phagosomal intensity was measured after subtracting cytosolic fluorescence. Data are expressed relative to the maximum value. Values in c and d are means ± SEM of three and five independent experiments respectively. Note the different time scales used in c vs. d. The initial contact between cells and IgG-SRBC was considered as time “0” throughout the figure.

cannot be attributed to resynthesis of PtdIns(4,5)P2, which remains undetectable.

We next compared PtdIns4P and PtdIns(3,4,5)P3 in cells co-transfected with mCh-2xP4M and

GFP-PH-Gab2 (Fig. 4.4a). As described earlier (Marshall et al., 2001), PtdIns(3,4,5)P3 is

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detectable in forming cups as targets are engaged and its levels increase as the pseudopods

extend. PtdIns(3,4,5)P3 persists on sealed phagosomes for ≈1 min and then disappears

abruptly. Neither the initial accumulation of PtdIns4P, nor its secondary disappearance appear

to be caused by degradation of or conversion to PtdIns(3,4,5)P3.

We also tracked PtdIns4P simultaneously with PtdIns3P, visualized using tandem FYVE

domains of EEA1. PtdIns3P appeared on the phagosomal membrane as PtdIns4P

disappeared, and was present throughout the period where PtdIns4P was absent (Figs. 4.4b

and d). Remarkably, this divergent behavior was also observed at the later stages, when

PtdIns4P re-appeared as PtdIns3P disappeared. While we are not aware of any mechanism

capable of directly interconverting these lipids, the changes in PtdIns4P and PtdIns3P seem to

be related, conceivably via signaling intermediates. Of note, based on extensive work

previously reported by us and others, we concluded that neither the phagocytic efficiency, the

time course of phagocytosis nor the changes in PtdIns(4,5)P2, PtdIns(3,4,5)P3 or PtdIns3P were

noticeably affected by the expression of the 2xP4M probe, confirming that it is innocuous at the

expression levels used in this study.

The reacquisition of PtdIns4P may represent a key—yet unappreciated—step in the transition

from early to late phagosomes. We therefore endeavored to place the PtdIns4P changes in the

context of defined stages of the maturation of the phagosome. To this end, we used different

maturation markers. Early phagosomes were identified by the presence of Rab5. As shown in

Fig. 4.5a, mRFP-Rab5 was acquired by the nascent phagosomes as PtdIns4P disappeared,

and was lost prior to the reacquisition of PtdIns4P. This inverse relationship resembles the

behavior described above for PtdIns3P, although this inositide persists longer on the early

phagosomal membrane than does Rab5, which ceases to be visible after 3-5 min (Fig. 4.5d).

Together, these data indicate that PtdIns4P is absent from the phagosome during the early

phase of maturation.

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Figure 4.4. Phosphoinositide metabolism during the early phagocytic stages. a) Time-lapse gallery of confocal micrographs of RAW 264.7 cells transiently co-expressing mCh-2xP4M and GFP-PH-Gab2 [a PtdIns(3,4,5)P3-specific biosensor] during phagocytosis of IgG-SRBCs. The Roman numerals refer to the stages defined on Figure 3. Images are representative of at least 10 similar determinations. b) Schematic diagram illustrating possible enzymatic reactions whereby phagosomes can be depleted of PtdIns4P. c) Confocal micrographs of RAW 264.7 cells co-expressing GFP-PH-TAPP1 [a PtdIns(3,4)P2

biosensor] and mCh-2xP4M during the phases of initial increase (left) and disappearance of PtdIns4P (right); PtdIns(3,4)P2 was not detected in phagosomes during PtdIns4P disappearance. d) Confocal micrographs acquired during phagocytosis of 1.6 µm TMR-IgG-coated latex beads by LY294002-treated (50 µM) RAW 264.7 cells expressing GFP-2xP4M. Representative images of the phases of initial PtdIns4P increase (left) and its subsequent disappearance (right) are shown. LY294002 did not prevent PtdIns4P disappearance. Insets show magnifications of dotted white boxes. e) RAW 264.7 cells were challenged with 3.87 µm IgG-coated latex beads. After 15 min, cells were washed and fixed, permeabilized and immunostained using anti-PI4KA antibodies. Confocal sections are shown. A forming phagosome is indicated with a yellow arrow while a maturing phagosome is indicated with a white arrow. Images are representative of at least 10 similar determinations. Scale bars = 5 µm.

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The reappearance of PtdIns4P coincides roughly with the onset of the late stage of maturation,

as defined by the acquisition of Rab7. Both GFP-2xP4M and mRFP-Rab7 are clearly visible on

phagosomes between 10-30 min after sealing occurred (Figs. 4.5b and d). The formation of

phagolysosomes follows shortly thereafter. Fusion of phagosomes with lysosomes was

assessed by pre-loading the latter with TMR-labeled dextran, using a well-established pulse-

chase protocol (see Methods). Maturing phagosomes fused with dextran-loaded lysosomes

slightly after PtdIns4P reacquisition (Fig. 4.5c). A comprehensive perspective of the temporal

relationship between the triphasic PtdIns4P changes relative to those of other phosphoinositides

and the canonical stages of phagocytosis is provided by the schematic cartoon in Fig. 4.5d.

4.3.3 Disappearance of PtdIns4P from the phagosome

Because conversion to multiphosphorylated species did not appear to account for the

disappearance of PtdIns4P from the nascent phagosome, we explored alternative mechanisms.

SAC2 (INPP5F) was recently characterized as a PtdIns 4-phosphatase that functions in Rab5-

positive early endosomes (Hsu et al., 2015; Nakatsu et al., 2015). Indeed, when co-expressed

with Rab5 in RAW 264.7 cells, GFP-SAC2 localized to endosomal structures. When analyzed

under similar conditions, GFP-SAC2 associated with early phagosomes when they sealed (Fig.

4.6a), at the same time as Rab5 did. Importantly, the association of SAC2 with phagosomes

coincided with the disappearance of PtdIns4P, monitored using 2xP4M (Fig. 4.6b).

We used gene silencing to assess whether this temporal coincidence reflected a causal

relationship, i.e. whether the acquisition of SAC2 was responsible for the loss of PtdIns4P. By

independent electroporation of two different small interfering RNA (siRNA) the expression of

SAC2 in RAW 264.7 cells was depressed ≈65%, as determined by qPCR (Fig. 4.6c). This

partial depletion of Sac2 resulted in a delayed disappearance of PtdIns4P from the formed

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Figure 4.5. Late phagosomes and phagolysosomes contain PtdIns4P. a) Time-lapse gallery of confocal micrographs of RAW 264.7 cells transiently co-expressing GFP-2xP4M and mRFP-Rab5 during phagocytosis of IgG-SRBCs. b) Confocal micrographs of RAW 264.7 cells transiently co-expressing GFP-2xP4M and mRFP-Rab7 during the course of phagocytosis of IgG-SRBCs. c) Confocal micrographs of cells expressing GFP-2xP4M and exposed to tetramethylrhodamine (TMR)-labeled 10 kDa dextran for 3 hrs and chased for 30 min to label lysosomes. Images in a-c are representative of the

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distribution noted during the indicated time intervals following the onset of phagocytosis; the insets show magnifications of the area delimited by dotted white boxes. Scale bars = 5 µm. d) Schematic diagram illustrating the relative timing of the acquisition by phagosomes of different phosphoinositides and of canonical markers of phagosome maturation.

phagosome (Figs. 4.6d and e) with both siRNA sequences. A fraction of the cells where SAC2

had been silenced did not entirely lose 2xP4M fluorescence from phagosomes even 3 min post-

internalization, contrasting with phagosomes from control cells (electroporated with non-

targeting siRNA), which consistently lost PtdIns4P ≈30-45 s after closure (Fig. 4.6c). While the

partial inhibition observed may be attributed to incomplete gene silencing, other mechanisms

could contribute to PtdIns4P reduction in the early phagosome.

We analyzed whether phospholipase C (PLC) activity contributes to the disappearance of

PtdIns4P from phagosomes. Depletion of PtdIns4P could conceivably result from its direct

hydrolysis by PLC (Zhang et al., 2013; Sicart et al., 2015) or may occur indirectly, following its

conversion to PtdIns(4,5)P2, the preferred substrate of the PLCs. Indeed, PLCγ1 and 2 are

primarily responsible for the disappearance of PtdIns(4,5)P2 during phagosome formation

(Azzoni et al., 1992; Liao et al., 1992; Scott et al., 2005). We therefore compared the rate of

formation of diacylglycerol (DAG), the product of hydrolysis of phosphoinositides by PLC, with

the disappearance of PtdIns4P. Because in macrophages DAG is rapidly converted to

phosphatidic acid (Bohdanowicz et al., 2013), to better visualize its accumulation we pre-treated

the cells with DAG kinase inhibitor II. In RAW 264.7 cells co-transfected with mCh-C1-PKC∂—

used as a probe for DAG (Shindo et al., 2003)—and GFP-2xP4M, DAG levels increased in

extending pseudopods as the phagosome sealed, at a stage when PtdIns4P underwent only a

modest increase (Figs. 4.7a and b). Maximal accumulation of DAG was observed as the

PtdIns4P decreased sharply, consistent with the notion that PLC activity contributed directly or

indirectly to the consumption of PtdIns4P. Accordingly, when recruited to the PM of otherwise

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Figure 4.6. Sac2 recruitment to phagosomes and PtdIns4P degradation. a) Confocal micrographs of RAW 264.7 cells co-expressing mCherry-2xP4M, GFP-Sac2 and CFP-Rab5 during phagocytosis of IgG-SRBCs (the CFP channel is not shown); insets show magnifications of the area delimited by dotted white boxes. Scale bar = 5 µm. b) Time course of the changes in PtdIns4P and Sac2 during phagosome formation, from experiments like the one illustrated in a) PtdIns4P was monitored using mCherry-2xP4M and normalized to plasmalemmal mCherry-2xP4M intensity (red line, black squares); GFP-Sac2 fluorescence in the phagosome was calculated after subtraction of GFP-Sac2 cytosolic intensity (green line, white squares); data are expressed relative to the maximum value. Values are means ± SEM from 3 independent experiments. In a) and b) pseudopod extension was considered as time “0”. c) Sac2 silencing efficiency of sequences siRNA1 and siRNA2 in RAW 264.7 cells measured by quantitative real-time PCR after reverse transcription; shown are the mean ± standard deviations of the mean of three independent experiments and normalized to cells treated with non-targeting siRNA (control). d) Time course of disappearance of PtdIns4P, assessed by kymography using GFP-2xP4M, during the early stages of phagocytosis. A schematic illustrating the region of the phagosome analyzed over time is shown at the top. The two lower panels show representative kymographs illustrating the disappearance

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of GFP-2xP4M from the base of the nascent phagosome over 120 sec. The closure of the phagosome was considered as time “0”. Images in a) and d) are representative of the distribution noted during the indicated time intervals following the onset of phagocytosis. e) GFP-2xP4M phagosomal intensity was measured and normalized to GFP-2xP4M plasmalemmal intensity; data are expressed relative to the maximum value. Blue lines illustrate the kinetics of GFP-2xP4M disappearance in cells treated with non-targeting (control) siRNA, red lines illustrate cells treated with Sac2-targetted siRNAs. White and black squares and associated bars show the means ± SE of control and Sac2-knockdown (using siRNA1 and siRNA2) cells, respectively, from at least 10 phagosomes from 3 independent experiments. The intersections of the dotted lines indicate the t1/2 values of decay. ★ P≤0.05, ★★P≤0.01, ★★★P≤0.005, ns = non significant.

resting cells using the rapamycin heterodimerization system, PLCβ3 (expressed as mRFP-

FKBP-PLCβ3) caused a marked reduction in PtdIns4P (Fig. 4.7d).

That PLC can directly hydrolyze PtdIns4P is suggested by the experiment illustrated in Fig. 4.7f.

In this instance mRFP-FKBP-PLCβ3 was recruited to the membrane of late phagosomes using

iRFP-FRB-Rab7 as the targeting determinant. As in the preceding case, addition of rapamycin

resulted in depletion of PtdIns4P. It is important to note that, unlike the PM, the late phagosome

contains PtdIns4P but no detectable PtdIns(4,5)P2. By analogy, it is conceivable that

plasmalemmal PtdIns4P may have undergone direct hydrolysis by PLC.

Finally, we considered whether a fraction of the PtdIns4P may have been converted to

PtdIns(3,4)P2 by class 2 phosphoinositide 3-kinases (PI3Ks; Fig. 4.4b). To investigate this

possibility, we examined the dynamics of PtdIns(3,4)P2 during phagocytosis using a biosensor

derived from the tandem-pleckstrin-homology-domain-containing protein (TAPP1), PH-TAPP1.

We found a modest, discrete accumulation of PtdIns(3,4)P2 in extending pseudopods as the

phagosome sealed (Fig. 4.4c), at a stage when PtdIns4P was undergoing accumulation.

However, PtdIns(3,4)P2 was no longer detectable when PtdIns4P underwent the acute depletion

that followed phagosome sealing. We also examined the disappearance of PtdIns4P from

forming phagosomes in the presence of the PI3K inhibitor LY294002. PI3K inhibition severely

impairs phagocytosis of large particles (>3-4 mm), but smaller ones are still ingested

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Figure 4.7. Assessment of the role of PLC in PtdIns4P disappearance. a) Confocal micrographs of RAW 264.7 cells co-expressing GFP-2xP4M and mCh-C1-PKC∂ (a DAG biosensor) during phagocytosis of IgG-SRBCs; cells were pre-treated with diacylglycerol kinase inhibitor II (30 µM) for 30 min to minimize the phosphorylation and rapid disappearance of DAG. b) Time course of the changes in PtdIns4P and DAG during phagosome formation. PtdIns4P was monitored using GFP-2xP4M and normalized to plasmalemmal GFP-2xP4M intensity (green line, white squares); mCh-C1-PKC∂ was used as DAG probe and normalized to mCh-C1-PKC∂ cytosolic intensity (red line, black squares). Data are expressed relative to the maximum value. Values are means ± SEM from 5 independent experiments. In a) and b) pseudopod extension was considered as time “0”. c) Schematic diagram illustrating the recruitment of PLCβ3 to the PM through a rapamycin heterodimerization system. d) RAW 264.7 cells transiently co-expressing GFP-2xP4M, mRFP-FKBP-PLCβ3 and Lyn11-FRB were imaged before (left) and after (right)

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the addition of rapamycin (1 µM). e) Schematic diagram illustrating the recruitment of PLCβ3 to Rab7-positive compartments (late phagosomes) through a rapamycin heterodimerization system. f. RAW 264.7 cells transiently co-expressing GFP-2xP4M, mRFP-FKBP-PLCβ3 and iRFP-FRB-Rab7 imaged before (left) and after (right) the addition of rapamycin; PLCβ3 recruitment caused release of GFP-2xP4M from the PM and from late phagosomes within seconds; insets show inverted images of mRFP-FKBP-PLCβ3 fluorescence. Scale bars = 5 µm.

(Cox et al., 1999a). We challenged cells pre-treated with LY294002 with 1.6 µm latex beads

and monitored PtdIns4P dynamics. As shown in Fig. 4.4d, inhibition of PI3Ks did not affect

PtdIns4P disappearance from small phagosomes upon their scission from the PM. Thus, we

found no evidence to suggest that phosphorylation by LY294002-sensitive PI3Ks contribute

measurably to the loss of PtdIns4P that follows phagosome sealing; similar results were

obtained using wortmannin (not shown). Instead, hydrolysis by SAC2, possibly aided by PLC

activity, are likely the major mechanisms accounting for the early disappearance of PtdIns4P.

4.3.4 PtdIns4P reappearance in maturing phagosomes

We then investigated the mechanisms underlying PtdIns4P reappearance in late phagosomes.

Mammalian cells express four different PtdIns4P kinases (PI4Ks): the class III enzymes

PI4KIIIα/PI4KA and PI4KIIIβ/PI4KB, and the class II enzymes PI4KIIα/PI4K2A and

PI4KIIβ/PI4K2B. (Balla, 2013; Boura and Nencka, 2015). PI4KA is responsible for maintaining

the plasmalemmal pool of PtdIns4P (Balla et al., 2008; Nakatsu et al., 2012), while both PI4KB

and PI4K2A function in the Golgi complex (Godi et al., 1999; Minogue et al., 2010). The class II

enzymes are thought to synthesize endo-lysosomal PtdIns4P (Balla et al., 2002; Salazar et al.,

2005; Jović et al., 2012).

To assess which of these enzymes is present on late phagosomes, we expressed fluorescently-

tagged chimeric constructs of each one of the PI4Ks in RAW 264.7 cells, and imaged them

during the course of phagocytosis of IgG-SRBCs. Only PI4K2A accumulated noticeably in

maturing phagosomes (Fig. 4.8a). PI4K2A recruitment was first apparent approximately 10 min

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Figure 4.8. PI4K2A recruitment and generation of PtdIns4P in maturing phagosomes. a) Confocal sections of RAW 264.7 cells expressing (i) GFP-PI4KA, (ii) GFP-PI4KB, (iii) GFP-PI4K2B, (iv) GFP-PI4K2A or (v) co-expressing GFP-PI4K2A and mCh-2xP4M. (vi) immunostaining of PI4K2A. Micrographs i - vi were acquired 20 min after initiation of phagocytosis. Insets show magnifications of the area delimited by dotted white boxes. b) Time course of the acquisition of PI4K2A and PtdIns4P during phagocytosis. PI4K2A was monitored measuring phagosomal GFP-PI4K2A and normalized to the mean fluorescence GFP-PI4K2A intensity of the entire cell (green line, white circles). Data are expressed relative to the maximum value. PtdIns4P was monitored measuring phagosomal mCh-2xP4M and normalized to plasmalemmal mCh-2xP4M (red line, black circles). Data are expressed relative to the value upon phagosomal closure. c) and d) PI4K2A silencing efficiency in RAW 264.7 (c) and COS-1-FcγRIIa cells (d), measured by quantitative real-time PCR after reverse transcription; results were normalized to control siRNA cells and are shown as means ± SD of at least 3 independent experiments. e) Confocal micrographs of COS-1-FcγRIIa cells expressing GFP-2xP4M treated with non-targeting

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(control) siRNA (left) and PI4K2A siRNA (right) after 40 min of phagocytosis. f) Fluorescence of phagosome-associated GFP-2xP4M normalized to plasmalemmal GFP-2xP4M in cells treated with non-targeting (control) siRNA (open bar) and PI4K2A siRNAs (black bars). Data are means ± SEM of four independent experiments of at least 23 phagosomes each. Red arrows indicate phagosomes. Scale bars = 5 µm.

after phagosome closure and the kinase was still clearly detectable after 30 min. When GFP-

PI4K2A was co-expressed with mCh-2xP4M, a correlation was observed between phagosomal

acquisition of the kinase and reappearance of PtdIns4P (Fig. 4.8av). Quantitation of multiple

experiments (Fig. 4.8b) showed that acquisition of the kinase slightly preceded the

reappearance of detectable amounts of the phosphoinositide in phagosomes. The recruitment

of PI4K2A suggested by the heterologous expression experiments was validated by

immunostaining. As illustrated in Fig. 4.8avi, association of the endogenous kinase with the late

phagosome was also clearly observed using the PI4K2A-specific mouse monoclonal antibody

4C5G.

To assess the importance of PI4K2A-mediated synthesis in the reappearance of PtdIns4P in

late phagosomes we used siRNA to silence the expression of Pi4k2a. We used various

combinations of six different oligonucleotides targeting the gene, but attained a maximum of

≈40% reduction in gene expression (e.g. Fig. 4.8c). Because gene silencing is notoriously

difficult in myeloid cells, we used an alternative approach. Balla et al. had reported

considerable success in silencing PI4K2A in COS cells (Balla et al., 2005), a green monkey

kidney cell line that is highly transfectable. While COS cells are not inherently phagocytic, we

and others had shown that they acquire the ability to engulf IgG-opsonized targets when

transfected with Fcγ receptors (Indik et al., 1991; Downey et al., 1999). Notably, the

phagosomes formed by such cells undergo maturation, acidification and acquire bacteriostatic

capacity (Downey et al., 1999). Indeed, COS-1-FcγRIIa cells –a line of COS-1 cells stably

expressing FcγRIIa– transiently expressing GFP-2xP4M and allowed to ingest IgG-SRBCs,

recapitulated the triphasic changes in phagosomal PtdIns4P described above for macrophages

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(Fig. 4.9). We proceeded to silence PI4K2A using an oligonucleotide sequence (siRNA1) for

validated previously in COS-7 cells (Wang et al., 2003; Balla et al., 2005), as well as a newly

designed sequence (siRNA2), to minimize the likelihood of off-target effects. Both sequences

caused effective (siRNA1≥80% and siRNA2≈70%) gene knockdown (Fig. 4.8d). Using this

approach, we evaluated the role of PI4K2A in phagosomes. As illustrated in Fig. 4.8e, while the

presence of PtdIns4P in the plasmalemma and the initial phase of loss from the phagosome

persisted, the late phase of reacquisition was markedly inhibited. The loss was somewhat

heterogeneous (GFP-2xP4M was virtually absent from phagosomes in some cells, while in

others the intensity of the biosensor was only partially reduced or unaffected), an observation

we attribute to the incomplete silencing of the PI4K2A gene. Nevertheless, the reduction of

phagosomal PtdIns4P was highly significant (p<0.0001; Fig. 4.8f), implying that PI4K2A is at

least partly responsible for the reformation of PtdIns4P in late phagosomes.

Figure 4.9. PtdIns4P dynamics in COS-1-FcγRIIa cells. a) Time-lapse gallery of confocal micrographs acquired during phagocytosis of TAMRA-labeled IgG-opsonized sheep red blood cells (IgG-SRBCs; shown in red) by COS-1-FcγRIIa cells transiently expressing GFP-2xP4M (green). The panels indicate the time elapsed from the moment the target was engaged. Insets show magnifications of the area

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delimited by dotted white boxes. b) Summary of the changes in PtdIns4P content of phagosomes following engagement of IgG-SRBCs. The intensity of GFP-2xP4M in phagosomes was quantified and normalized to that of the plasmalemma. The three phases defined in RAW 264.7 were recapitulated in COS-1-FcγRIIa cells, with slower dynamics. Data are expressed relative to the maximum value attained during the initial transient increase, which was defined as time “0”. Values are means ± SEM of 4 independent experiments. Scale bars = 5 µm.

4.3.5 PtdIns4P is required for completion of phagosome maturation

The synchronous disappearance of PtdIns3P and resurgence of PtdIns4P coincide with the

early-to-late transition of phagosomes. The phosphoinositide switch could be the cause or a

consequence of the maturation process. This was analyzed by impairing the formation of

PtdIns4P through silencing of PI4K2A, and analyzing the effects of this inhibition on phagosome

maturation. The early-to-late transition is characterized by a marked drop in luminal pH, as the

phagosomes acquire V-ATPases. We therefore compared the accumulation of an acidotropic

fluorescent dye, an index of acidification, in phagosomes of cells treated with non-targeting or

PI4K2A-targetting siRNA. We used cresyl violet, a fluorescent weak base that accumulates

within acidic organelles (Fig. 4.10a) (Ostrowski et al., 2016). Its suitability as an indicator of

acidic pH was confirmed by its accumulation in lysosomes, which were identified by pre-loading

with labeled dextran using a well-established pulse and chase protocol (Fig. 4.10b). Moreover,

cresyl violet co-localized extensively with GFP-Rab7, while no significant co-localization was

observed with GFP-Rab5 (Fig. 4.11), validating its use as a marker of late endocytic

compartments. When analyzed 35-40 min after particle internalization, phagosomes of cells

treated with non-targeting (control) siRNA regularly acquired cresyl violet (Fig. 4.10c left).

Strikingly, cresyl violet was not detectable in most phagosomes of cells where PtdIns4P

reappearance was impaired using PI4K2A siRNA (Fig 4.10c, right). Interestingly, cresyl violet-

positive vesicles were often observed surrounding such phagosomes, suggesting that fusion,

rather than the acidification of lysosomes, was impaired when PI4K2A was silenced. Because

not all cells/phagosomes were equally affected by the siRNA treatment, we quantified both the

amount of PtdIns4P remaining on phagosomes (relative to that on the PM) and the extent to

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Figure 4.10. Phagosome acidification is impaired when PI4K2A is silenced. a) Schematic illustrating the structure of cresyl violet and the proposed mechanism whereby it accumulates in acidic compartments; note protonation of cresyl violet occurring in dotted red box. b) Single confocal section of

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COS-1-FcγRIIa cells where lysosomes were loaded with Alexa fluor-647 10 kDa dextran (0.1 mg/mL, 3-h pulse, 30-min chase) followed by cresyl violet loading (1 µM, 2-min pulse); insets show magnifications of the area limited by the dotted line. c) Confocal micrographs of cells treated with non-targeting (control) siRNA (left) and PI4K2A siRNA (right) COS-1-FcγRIIa cells expressing GFP-2xP4M that were pulsed with cresyl violet after 40 min of phagocytosis of IgG-SRBC. Scale bars = 5 µm. d) Plot relating cresyl violet acquisition with PtdIns4P levels in phagosomes, measured 40 min after phagocytosis; r2 = 0.64. The vertical red line represents an arbitrary threshold dividing two phagosomal populations based on phagosomal GFP-2xP4M intensity relative to that in the plasma membrane. White squares represent phagosomes with low PtdIns4P levels, black squares represent phagosomes with higher PtdIns4P levels.

which they accumulated the acidotropic dye. Data obtained from 89 phagosomes in four

separate experiments are collated in Fig. 4.10d. There is a clear correlation (r2 = 0.64) between

these parameters, strongly suggesting that accumulation of PtdIns4P by late phagosomes is

essential for their full acidification.

Because prolonged and generalized absence of PI4K2A may have affected other cellular

compartments, potentially causing indirect effects, we used an additional approach to assess

the role of PtdIns4P in phagosome maturation. To circumvent such potentially non-specific

Figure 4.11. Cresyl violet co-localizes with Rab7, but not with Rab5. A) Representative confocal section of RAW 264.7 cells transiently expressing GFP-Rab5 and treated with cresyl violet (1 µM for 2 min; shown in red). B) Single confocal section of RAW 264.7 cells transiently expressing GFP-Rab7 treated with cresyl violet as above. Insets show magnifications of dotted white boxes for each channel. Scale bars = 5 µm.

effects, we acutely depleted phagosomal PtdIns4P by recruiting a PtdIns4P-specific

phosphatase, SAC1, using rapamycin-mediated heterodimerization. mCh-FKBP-SAC1 was

recruited to Rab7-containing (late) phagosomes by co-expression with iRFP-FRB-Rab7 (Fig.

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4.12a). The cells were also co-transfected with GFP-2xP4M to monitor the effects of

phosphatase recruitment on PtdIns4P. Addition of rapamycin caused depletion of PtdIns4P

from phagosomes that had formed ≥35 min earlier (Fig. 4.12b). Using this paradigm, we

proceeded to study the association of the lysosomal-associated protein RILP –an effector of

active Rab7– with phagosomes. We used a chimeric construct that encodes for, the C-terminal

domain RILP (Cantalupo et al., 2001); in control cells GFP-RILP-C33 decorates the entire

phagosomal membrane, consistent with extensive and sustained activation of Rab7 (Figs. 4.12c

and d). Strikingly, in cells where PtdIns4P was depleted by recruitment of SAC1, GFP-RILP-

C33 acquisition by phagosomes was impaired compared to control cells (expressing mCh-FKBP

not attached to SAC1) (Figs. 4.12c and d). Note that in mCh-FKBP-SAC1-transfected cells, the

construct was recruited normally to the phagosomes, indicating that the localization of iRFP-

FRB-Rab7 was not affected. The targeting of iRFP-FRB-Rab7 to the membrane seemingly

does not require activation of the GTPase. Additionally, we observed that GFP-RILP-C33-

positive vesicles often accumulated around PtdIns4P-depleted phagosomes, consistent with the

earlier suggestion that fusion with late-endosomes/lysosomes was impaired. Time-lapse video

imaging supported the notion that incoming vesicles failed to fuse with phagosomes, and

showed no evidence that the observed vesicles were budding-off the phagosomal membrane.

Lastly, when mRFP-Rab7 was transiently co-expressed with GFP-RILP-C33 in PI4K2A-silenced

cells, a similar effect was observed. Rab7 accumulated in the phagosomal membrane, while

RILP-C33 recruitment was generally impaired when compared to control cells (Fig. 4.12e).

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Figure 4.12. RILP acquisition is impaired in PtdIns4P-depleted phagosomes. a) Schematic diagram illustrating the recruitment of Sac1 to the PM through a rapamycin heterodimerization system. b) COS-1-FcγRIIa cells transiently co-expressing GFP-2xP4M, mCh-FKBP-Sac1 and iRFP-FRB-Rab7 were challenged with IgG-SRBCs; phagocytosis was allowed to develop for 30 min before imaging. Upon

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PtdIns4P reappearance in phagosomes, cells were treated with rapamycin (1 µM) to induce recruitment of mCh-FKBP-Sac1, which resulted in PtdIns4P hydrolysis (release of GFP-2xP4M from phagosomes). Insets show mCh-FKBP-Sac1 fluorescence (black and white were inverted for clarity). c) and d) COS-1-FcγRIIa cells transiently co-expressing GFP-RILP-C33, mCh-FKBP-Sac1 and iRFP-FRB-Rab7; in control cells mCh-FKBP-Sac1 was substituted with mCh-FKBP. The cells were challenged with IgG-SRBCs, treated with rapamycin (1 µM) and phagocytosis was allowed to proceed for 40 min. Cells were then washed, fixed and imaged. d) Shows representative confocal micrographs of mCh-FKBP- (left) and mCh-FKBP-Sac1-expressing cells (right); insets show magnifications of the dotted white boxes for each channel. c) Shows GFP-RILP-C33 accumulation in phagosomes of cells expressing mCh-FKBP (open bars) or mCh-FKBP-Sac1 (black bars); phagosomes showing a continuous pattern around the phagosome, and those with only punctate or no fluorescence were scored separately. e) Confocal micrographs of COS-1-FcγRIIa cells co-expressing mRFP-Rab7 and GFP-RILP-C33 treated with non-targeting (control) siRNA (left) and PI4K2A siRNA (right) after 40 min of phagocytosis; insets show magnifications of the dotted white boxes. Scale bars = 5 µm.

4.4 Discussion

We observed localized triphasic changes in the level of PtdIns4P during phagocytosis. These

are summarized in schematic form in Fig. 4.13. Initially, PtdIns4P accumulated in the forming

phagocytic cup. This coincided with the increase in PtdIns(4,5)P2 in extending pseudopods,

which had been reported earlier (Botelho et al., 2000). The accumulation of PtdIns4P in this

setting may reflect localized synthesis required to satisfy the enhanced substrate demand for

stimulated PtdIns(4,5)P2 generation. In this regard, we found a discrete accumulation of

endogenous PI4KIIIα (PI4KA) in forming phagosomes (Fig. 4.4a). This raises the intriguing

possibility that the PI4KIIIα complex (PI4KA-TTC7-EFR3) responsible for the plasmalemmal

pool of PtdIns4P (Wu et al., 2014; Chung et al., 2015a)– may undergo stimulation at the cup.

Upon phagosome closure, PtdIns4P reaches a peak that coincides with the sudden

disappearance of PtdIns(4,5)P2 from the vacuolar membrane. We believe that these events are

linked in two ways. First, a previous study showed that the 5-phosphatases OCRL and INPP5B

are recruited to nascent phagosomes (Bohdanowicz et al., 2012b). These enzymes

dephosphorylate PtdIns(4,5)P2, yielding PtdIns4P. In addition, cessation of PtdIns(4,5)P2

synthesis likely contributes to the accumulation of PtdIns4P. PIP5Ks that use PtdIns4P to

synthesize PtdIns(4,5)P2 localize to the PM and are present at the phagocytic cup. However,

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these kinases are acutely released from the membrane of phagosomes as they undergo closure

(Fairn et al., 2009). Detachment of the PIP5Ks while PtdIns4P production is ongoing would

result in a localized burst of PtdIns4P. The observed transient accumulation of PtdIns4P likely

results from the combination of the above-mentioned phenomena.

Figure 4.13. Diagrammatic representation of the changes undergone by phosphoinositides during phagosome formation and maturation. The changes in PtdIns4P (shades of blue) are compared with those undergone by PtdIns(4,5)P2 (green), PtdIns(3,4,5)P3 (red) and PtdIns3P (purple). Phagosome formation is divided into receptor engagement, pseudopod extension and vacuole sealing stages, while maturation is divided into early and late/lysosomal stages.

The peak noted upon sealing is followed within 30-45 s by the virtual disappearance of

PtdIns4P, which is undetectable in the membrane of early phagosomes for a period lasting ≈10

min (Figs. 4.1f, g and 4.4d). Our data suggest that SAC2 mediates at least part of the

hydrolysis of PtdIns4P, but PLC activity also seems to contribute, whether by depleting

PtdIns(4,5)P2 and thereby accelerating PtdIns4P consumption, or by directly hydrolyzing the

latter (Figs. 4.6 and 4.7). Exchange of PtdIns4P for other lipids (e.g. cholesterol) via contacts

with the endoplasmic reticulum may have also occurred, but was not investigated here. A

period of about 10 min when PtdIns4P is undetectable follows its abrupt disappearance after

sealing. Strikingly, PtdIns4P then reappears. This secondary accumulation of PtdIns4P

reaches its maximum ≈30 min after the onset of phagocytosis, attaining levels that clearly

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exceed those detected in the PM. The late reappearance of PtdIns4P coincides with and

depends on the recruitment of PI4K2A.

Remarkably, the multiphasic changes in PtdIns4P are accompanied by opposite changes in the

concentration of phagosomal PtdIns3P. The initial loss of PtdIns4P coincides with the

emergence of PtdIns3P, and the converse is true once the early maturation stage is completed:

PtdIns3P disappears abruptly as the phagosome regains PtdIns4P (Figs. 4.3, 4.5 and 4.13). A

similar phospholipid conversion process was recently described to occur at the sorting-to-

recycling endosome interphase (Ketel et al., 2016). Thus, the identity and function of the early

endosomal and phagosomal compartments may require not only acquisition of PtdIns3P, but

also the simultaneous elimination of PtdIns4P. In this regard, it is noteworthy that the class III

PI3K Vps34 that synthesizes PtdIns3P and the 4-phosphatase SAC2 co-exist in early

endosomes/phagosomes (Hsu et al., 2015; Nakatsu et al., 2015), while the PtdIns4P kinase

PI4K2A and some myotubularins –3-phosphatases that break down PtdIns3P– localize to the

late compartments (Tsujita et al., 2004; Lorenzo et al., 2006). These observations are

compatible with the suggestion of Ketel and colleagues that activation of the myotubularins that

eliminate PtdIns3P may be coupled to the acquisition of the kinases that foster PtdIns4P

formation (Ketel et al., 2016). We suggest that PtdIns4P, possibly in conjunction with

PtdIns(3,5)P2, defines the identity and functional properties of the late

phagosome/phagolysosome. This speculation is in accordance with recent reports showing that

PtdIns4P plays a role in the fusion of autophagic vacuoles with lysosomes (Wang et al., 2015a)

and possibly also in phagolysosome biogenesis, as suggested from heterologous fusion

experiments in cell-free systems (Jeschke et al., 2015).

How PtdIns4P regulates late maturation stages remains unclear. The only known PtdIns4P-

specific effectors, such as the ceramide transfer protein CERT, the glucosylceramide transfer

protein FAPP2 and the oxysterol binding protein OSBP1 function at the Golgi level (Graham and

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Burd, 2011). Our observations indicate that proper acidification requires the build-up of

PtdIns4P in late phagosomes, an effect that may be mediated by RILP (Figs. 4.10 and 4.12).

Indeed, interference with RILP function was shown earlier to impair phagolysosome formation

(Harrison et al., 2003). RILP forms a complex with ORP1L that affixes dynein to Rab7-

containing compartments (Johansson et al., 2007) and drives the centripetal motion of

phagosomes towards the juxtanuclear region where lysosomes reside. It is of interest that

ORP1L possesses a pleckstrin-homology domain known to bind phosphoinositides (Johansson

et al., 2005). We speculate that PtdIns4P is required for the recruitment of effectors to maturing

phagosomes. Potential effectors include the V-ATPases, which recruitment to the Golgi

apparatus in yeast was recently shown to be dependent on PtdIns4P (Banarjee and Kane,

2017).

In summary we have defined the dynamics of PtdIns4P during phagocytosis in live

macrophages, and have begun to elucidate the underlying mechanisms behind these changes

and their functional implications. The phosphoinositide clearly undergoes pronounced changes

and is likely to play additional, unappreciated roles in phagosome biology. The mechanism

whereby PI4K2A is recruited to maturing phagosomes and the possible role of PtdIns4P in

phagosome resolution are of particular interest and should be the focus of future studies.

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Chapter 5 Phosphatidylinositol 4-phosphate Regulation by Endoplasmic

Reticulum – Phagolysosome Contact Sites Directs Phagosome Resolution

This chapter has been modified from the following submitted manuscript: Roni Levin, Fernando

Montaño, Tal Keren-Kaplan, Braeden Ego, Jessica Diciccio, William S. Trimble, Michael C.

Bassik, Juan Bonifacino, Gregory D. Fairn and Sergio Grinstein.

Phosphatidylinositol 4-phosphate regulation by 5endoplasmic reticulum – phagolysosome contact sites directs phagosome resolution

5.1 Abstract

Conceptually, phagocytosis can be divided into three stages: phagosome formation, maturation

and resolution. Tight coordination of the compendium of events that span these three stages

requires precise regulation of phosphoinositide metabolism. Indeed, these phospholipids

undergo marked acute changes during phagocytosis. These changes have paramount

physiological implications in the progression of the endocytic process. While phosphoinositides

have been widely characterized during the early stages phagocytosis, little is known about these

them during the late maturation and resolution stages. Recently, we defined the presence of

PtdIns4P as an important player during the late maturation stage. Here we identified ORP1L—a

Rab7 effector—as a PtdIns4P-transfer protein. ORP1L transported PtdIns4P from the

phagolysosome to the ER while tethering both compartments. This resulted in a discontinuous

PtdIns4P distribution along the phagosomal membrane where the lipid was enriched in confined

regions and depleted from others. From these PtdIns4P-rich regions, we identified tubular

structures and fission events marking the initiation of phagosome resolution. Additionally, these

structures were enriched in Arl8b and SKIP, and accumulation of the latter was PtdIns4P-

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dependent. Finally, disruption of PtdIns4P metabolism during late stages impaired phagosome

resolution.

5.2 Introduction

Phagocytosis is an essential component of the innate immune response. Professional

phagocytes, such as macrophages, neutrophils and dendritic cells have the ability to internalize,

degrade and dispose of pathogenic microorganisms (Stuart and Ezekowitz, 2005; Flannagan et

al., 2012; Levin et al., 2016). Additionally, phagocytosis is central for tissue homeostasis:

jointly, professional and non-professional phagocytes clear in excess of 200 billion effete cells

per day from the human body (Elliott and Ravichandran, 2010; Bianconi et al., 2013; Elliott and

Ravichandran, 2016).

Phagosome formation is initiated by receptor engagement and clustering, followed by actin

rearrangement and scission of the nascent vacuole from the plasma membrane (PM)

(Swanson, 2008; Flannagan et al., 2012; Levin et al., 2016). Once formed, the phagosome

undergoes rapid and intense remodeling through interaction with components of the endocytic

pathway. During this transition—known as phagosome maturation—the phagosome acquires

microbicidal and degradative properties (Underhill, 2005; Kinchen and Ravichandran, 2008;

Fairn and Grinstein, 2012). Both phagosome formation and maturation have been studied in

extensive detail. By contrast, much less is known about the concluding stage, namely

phagosome resolution, whereby phagolysosomes and their contents are resorbed. Resolution

entails elimination and often re-utilization of the phagosomal contents, and recycling of cellular

components in preparation for additional rounds of phagocytosis (Levin et al., 2016; Gray and

Botelho, 2017). Antigen presentation to lymphoid cells is also a consequence of phagosome

resolution (Blander and Medzhitov, 2006; Mantegazza et al., 2013; Levin et al., 2016). Despite

its obvious importance, the molecular basis of phagosome resolution has not been explored and

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our current view of the process is limited to speculation and extrapolation from processes such

as lysosome turnover and autophagy.

Phagosome formation and maturation are associated with marked, acute changes in

phosphoinositide metabolism (Gillooly et al., 2001; Deretic et al., 2007; Levin et al., 2015).

Formation of the phagosome is accompanied by focal generation of PtdIns(3,4,5)P3 and

disappearance of PtdIns(4,5)P2 (Araki et al., 1996; Botelho et al., 2000; Marshall et al., 2001),

which are required for actin restructuring, pseudopod extension and vacuolar sealing (Araki et

al., 1996; Cox et al., 1999a; Rohatgi et al., 2000b). The newly formed PtdIns(3,4,5)P3

disappears promptly from the sealed phagosome, where it is followed immediately by a burst of

PtdIns3P formation (Fratti et al., 2001a; Vieira et al., 2001). The latter persists for several

minutes, coinciding with the early stages of the maturation sequence. Unlike these

phosphoinositides, PtdIns4P undergoes multiphasic changes during the phagosome formation

and maturation stages: its concentration, which is sizeable in the PM, increases in the PM upon

scission, followed by an abrupt decline. PtdIns4P is undetectable during the early maturation

stages, but reappears when PtdIns3P disappears, reaching a concentration that is noticeably

higher than that of the PM (Levin et al., 2017).

While the roles of other phosphoinositides in phagosome formation and maturation have been

studied and at least partially elucidated, little is known about the function of PtdIns4P; to date,

there are no known PtdIns4P effectors associated with the phagosomal membrane. Because it

is present and has functional implications at the end of the phagosome maturation sequence

(Jeschke et al., 2015; Levin et al., 2017) and since it was preliminarily detected in tubular

structures emanating from phagolysosomes (Levin et al., 2017), we speculated that PtdIns4P

might play a role in phagosome resolution. Here we describe yet another phase in the

metabolic conversion of PtdIns4P during phagocytosis, the underlying mechanism and its

relationship to the events that initiate phagosome resolution.

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5.3 Results

5.3.1 Phagosome resolution

Phagosome fission events have been reported previously (Krajcovic et al., 2013), but the

sequence of events that lead to the resolution of the phagolysosome has not been studied in

detail. We therefore initially documented the progressive fragmentation of phagosomes formed

by RAW 264.7 macrophages that had ingested IgG-opsonized sheep erythrocytes (IgG-SRBC).

To visualize the outline of the phagosomes, the macrophages were transfected with soluble

(cytosolic) GFP, while the SRBC were covalently labeled with tetramethylrhodamine (TAMRA in

Fig. 5.1a). Based on the size of the phagosomes and the integrity of their cargo, we identified

three major stages of late maturation, all occurring after fusion with lysosomes had taken place

(i.e. >20 min after RBC engulfment): stage I, which was predominant at the early stages of

phagolysosome formation, consisted of homogeneously sized (3 – 5 µm) vacuoles containing

seemingly intact, undigested SRBC; stage II consisted of largely intact phagosomes that

contained partially digested SRBC, with small fragmented structures in close proximity; in stage

III, the phagocytic vacuoles were mostly fragmented into smaller vesicles dispersed throughout

the cytoplasm, containing SRBC remnants (Fig. 5.1a). As shown in Fig. 5.1c resolution

occurred progressively, with ≈70% of the phagosomes having fragmented (stage III) after ≈3

hours. We gained additional insight of the structures formed at the individual stages by

analyzing samples at varying times after phagocytosis by electron microscopy (Fig 5.1b).

These experiments confirmed that stage I phagolysosomes contained mostly intact SRBC, and

that some of the SRBC had undergone lysis by stage II, evident by the appearance of areas of

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Figure 5.1. Phagosomal resolution is characterized by tubular fission events. a) Confocal slices of GFP-expressing RAW macrophages after phagocytosis of TAMRA-labeled SRBC; micrographs are representative of three stages of phagosomal resolution defined based on the integrity of the target and of the phagosome; scale bars = 10 µm. b) Representative transmission electron micrographs of RAW macrophages during the three stages of resolution of SRBC-containing phagosomes; for stage III, SRBC were pre-labeled with ferritin; scale bars = 200 nm. c) Assessment of phagosome resolution in RAW macrophages over time. d) Confocal slices representative of membrane remodeling in phagolysosomes of RAW macrophages expressing mCh-LAMP1; scale bars = 5 µm. e) Representative scanning electron micrographs of structures budding and extruding from resolving phagosomes with either unlabeled (left) or ferritin-labeled (right) SRBC; scale bars = 200 nm.

reduced electron density indicative of hemoglobin leakage. Hemoglobin-rich vesicles and

tubules were also seen in the vicinity of the phagosomes at this stage. Multiple smaller vesicles

of intermediate electron density were seen at stage III. That these derived from the

fragmentation of phagosomes was verified by labeling the SRBC with cationized ferritin prior to

phagocytosis. As shown in the rightmost panel of Fig. 5.1b, ferritin was readily discernible in the

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vesicles. Phagosome fragmentation was further confirmed by the use of a LAMP1 construct

tagged with a photoactivatable GFP (LAMP1-PAGFP). We assessed vesiculation by

exclusively photoactivating the phagosomal membrane of phagolysosomes (Fig. 5.2b) at the

early stages of phagolysosome biogenesis and following the distribution of PAGFP-positive

membranes over time. As shown in Fig. 5.2c, LAMP1-PAGFP-containing vesicles derived from

the phagosome appeared during maturation.

The dynamic nature of the remodeling process was visualized by expressing fluorescently

tagged LAMP1. Using confocal microscopy, we initially detected fusion of LAMP1-bearing

vesicles with the maturing phagosomes, followed by the extension of tubular protrusions from

the mature phagolysosomes, some of which underwent fission or formed blebs that remained

associated with the principal vacuole for varying periods (Fig. 5.1d). Most often, areas of the

phagosomal membrane where LAMP1 appeared thicker were noted. Because these were not

well resolved by optical microscopy, we analyzed their structure by electron microscopy. As

shown in Fig. 5.1e, they consisted of intricate tubular networks identified as connected to the

phagosome by the electron density of hemoglobin (e.g. bottom left) and/or of ferritin (bottom

right).

5.3.2 PtdIns4P dynamics during the early stages of phagosome resolution

Because phagolysosome formation is associated with reacquisition of PtdIns4P (Jeschke et al.,

2015; Levin et al., 2017), we speculated that this inositide may play a role in the remodeling and

resolution sequence. As shown in Figs. 5.3a-b, the highest concentration of PtdIns4P,

visualized using the 2xP4M biosensor (Hammond et al., 2014), was attained 20-30 min after

particle internalization, the time when resolution of the phagolysosomes begins. After reaching

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Figure 5.2. Dispersed compartments during phagosome resolution are of phagosomal origin. a) Transmission electron micrographs of RAW macrophages during stage II of phagosome resolution of IgG-SRBC pre-labeled with ferritin. Scale bars+ 10 µm. b) Confocal micrograph of RAW macrophages after 30 minutes of phagocytosis of IgG-SRBC (blue). Cells were expressing LAMP1-PAGFP. Micrographs were acquired before (left) and after activation of LAMP1-PAGFP in the membrane of an individual phagosome (right). c) Time-lapse micrographs of RAW macrophages expressing LAMP1-PAGFP during the three stages of phagosome resolution; the phagosomal membrane was photo-activated after 30 min of phagocytosis. Scale bars = 10 µm.

peak levels, the phosphoinositide gradually disappeared over the next 45 min (Fig. 5.3a-b).

This decrease was characterized by preferential depletion of PtdIns4P in discrete regions of the

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phagosomal membrane, with enrichment in the adjacent regions (Fig. 5.3a). More notably, we

observed PtdIns4P-positive vesicular and tubular structures emanating from the areas of the

phagosome where the phosphoinositide was enriched (Fig. 5.3c). To obtain better spatial and

temporal resolution of these events, we turned to lattice light-sheet microscopy. The three-

dimensional images acquired by this method showed striking tubulation events (Fig. 5.3d).

Although the ultimate destination of these tubules could not be ascertained, a fraction of them

approached the Golgi complex while others seemed directed to the plasma membrane.

5.3.3 PtdIns4P degradation mechanism

We next investigated the mechanism underlying the inhomogeneous distribution of PtdIns4P

during its late disappearance stage. First, we considered the possibility that PI4K2A—the

kinase shown earlier to be responsible for the reappearance of PtdIns4P in phagolysosomes

(Jeschke et al., 2015; Levin et al., 2017)—might redistribute and ultimately detach from the

membrane. However, we found the kinase to remain associated with the phagolysosomal

membrane throughout the period when PtdIns4P was depleted (Fig. 5.4). These observations

suggested that variations in the rate and location of PtdIns4P removal were responsible for its

inhomogeneous depletion. PtdIns4P depletion could in principle be catalyzed by lipid

phosphatases or phospholipases or by kinases that could convert it to other species. To our

knowledge, no such enzymes with selectivity for PtdIns4P have been described in late

endosomes, lysosomes or late phagosomes. Hence, we turned our attention to a potential lipid-

transport mechanism, namely oxysterol-binding protein-related protein 1L (ORP1L). ORP1L

contains a series of ankyrin repeats that bind Rab7 (Johansson et al., 2005) and is therefore

anticipated to associate with phagolysosomes, which are Rab7-positive. In addition, ORP1L

has a pleckstrin homology (PH) domain that binds phosphoinositides with low affinity and

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Figure 5.3. Dynamics and distribution of PtdIns4P during initiation of phagosome resolution. a) Top: time-lapse gallery of confocal micrographs acquired during phagocytosis of SRBC by RAW macrophages transiently expressing GFP-2xP4M showing multiphasic PtdIns4P dynamics (top); bottom: time-lapse surface intensity plots corresponding to the phagosome in the insets above; scale bar = 10 µm. b) Summary of tetra-phasic changes of PtdIns4P during phagocytosis; for quantitation, phagosomal fluorescence intensity was normalized to plasmalemmal fluorescence intensity; shown are means of 10 experimental replicates, error bars represent SEM c) Confocal micrographs showing phagosomal tubules decorated with GFP-2xP4M. d) Top: side view of lattice light-sheet time-lapse micrographs of RAW macrophages expressing GFP-2xP4M containing phagolysosomes; bottom: 3D surface reconstructions of GFP-2xP4M-positive phagosomes during tubulation for resolution. 3D reconstructions correspond to LLS micrographs on top; green surfaces are in contact with the phagosome and yellow surfaces are other compartments. Top scale bars = 10 µm; bottom scale bars = 3 µm.

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Figure 5.4. PI4K2A remains in the phagosomal membrane during maturation and initiation of phagosome resolution. Representative confocal micrographs of RAW macrophages expressing GFP-PI4K2A during the transition from late phagosome maturation to early phagosome resolution. a) 45 min; b) 60 min; c) 80 min. Scale bars = 10 µm.

specificity (Johansson et al., 2005), an FFAT motif that can associate with the ER proteins

VAPA/B (Loewen et al., 2003; Loewen and Levine, 2005; Rocha et al., 2009) and an OSBP-

related domain (ORD) shown to bind and translocate sterols (Suchanek et al., 2007; Rocha et

al., 2009; Vihervaara et al., 2011; van der Kant et al., 2013a; Wijdeven et al., 2016; Zhao and

Ridgway, 2017) (Fig. 5.5a). Of note, while the ORD domain of ORP1L can indeed bind

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cholesterol, it was unexpectedly found to bind PtdIns4P with even higher affinity (Zhao and

Ridgway, 2017). Because SAC1—the main 4-phosphatase of mammalian cells (Balla, 2013)—

localizes primarily to the ER (Rohde et al., 2003), we hypothesized that ORP1L might mediate

PtdIns4P transfer from the phagolysosome to the ER, where it would be degraded by SAC1

(Zewe et al., 2018). Analogous lipid transport mechanisms have been reported in other

compartments (Mesmin et al., 2013; Chung et al., 2015b; von Filseck et al., 2015; Moser von

Filseck et al., 2015; Sohn et al., 2018). To assess this hypothesis we first analyzed whether

ORP1L was expressed in myeloid cells. We found that, consistent with earlier transcriptomic

data (Johansson et al., 2003), ORP1L protein is present in monocytes and is markedly

upregulated upon differentiation to macrophages (Fig. 5.5b). Notably, expression of the short

variant of ORP1 (ORP1S) was particularly low (Fig. 5.6). In primary macrophages, endogenous

ORP1L localized to endomembranes, as did GFP-tagged ORP1L expressed heterologously in

RAW macrophages (Fig. 5.5c). The endomembrane compartment corresponded to late

endosomes/lysosomes, as it was Rab7-positive (Fig. 5.5d). That interaction of its ankyrin

repeats with Rab7 was responsible for ORP1L association with endo/lysosomes was verified by

expressing Rab7(T22N), a dominant-negative form of the GTPase, which resulted in

displacement of ORP1L to the cytosol (Fig. 5.5e). In primary macrophages that had ingested

opsonized targets, ORP1L decorated the phagolysosomal membrane (Fig. 5.5f), as did GFP-

tagged ORP1L in RAW macrophages (Fig. 5.5g). ORP1L was first detected on phagosomes

about 15 min after their formation and attained maximal levels after ≈30 min (Fig. 5.5h),

paralleling the course of acquisition of Rab7. It is noteworthy that while GFP-ORP1L initially

associated with phagosomes in a discontinuous, “patchy” pattern that resembled that of the

endogenous protein, the continued recruitment of the ectopically (over)expressed protein

gradually made the pattern more continuous. The rate and extent of this transition were

proportional to the level of expression of GFP- ORP1L.

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Figure 5.5. ORP1L accumulates in phagosomes during late maturation. a) Schematic representation of functional domains within ORP1L. b) Representative western blot showing ORP1L expression in primary human monocytes and macrophages, and in murine RAW macrophages. c) Endomembrane localization of endogenous ORP1L in primary human macrophages (left); endomembrane localization of expressed GFP-ORP1L in RAW macrophages (right). d) Confocal

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micrographs of RAW macrophages co-expressing GFP-ORP1L and RFP-Rab7 in RAW macrophages; insets are magnifications of red boxes; yellow lines show the cell periphery. e) Confocal micrographs of RAW macrophages co-expressing GFP-ORP1L and dominant-negative RFP-Rab7(T22N). f) Endogenous localization of ORP1L in primary human macrophages during phagocytosis of IgG-opsonized human RBC; yellow lines show the cell periphery (left); magnification of phagosome within red box (right). g) RAW macrophage expressing GFP-ORP1L during phagocytosis; yellow lines show the cell periphery (left); magnification of phagosome within red box (right). h) Kinetics of accumulation of ORP1L in maturing phagosomes of RAW macrophages; phagosomal ORP1L intensity was normalized to ORP1L intensity in the rest of the cell; shown are means of five experimental replicates, error bars represent SEM. Scale bars = 10 µm throughout the figure.

Figure 5.6. Monocytes and macrophages express low levels of ORP1S. The polyclonal ORP1 antibody recognizes the C-terminus of ORP1L and ORP1S. Lane 1: non-differentiated primary human monocytes; lane 2: MCSF-differentiated human macrophages; lane 3: PMA-treated U937 cells; lane 4: RAW macrophages. Anti-GAPDH was used as a loading control.

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Having validated the association of ORP1L with phagolysosomes, we proceeded to test whether

the protein plays a role in the late, progressive disappearance of PtdIns4P described above. To

this end, we co-expressed 2xP4M and ORP1L in RAW macrophages. Strikingly, we observed

that the ORP1L-rich regions of the phagosomes were preferentially depleted of PtdIns4P (Figs.

5.7a and 5.8). The mutual exclusion was exacerbated over time, as ORP1L levels gradually

increased in the phagosome. Eventually ORP1L accumulated throughout most of the

phagosome, at which point PtdIns4P was virtually undetectable (Fig. 5.7a).

5.3.4 ORP1L-dependent phagolysosome-to-ER PtdIns4P transport

The preceding observations suggest a role for ORP1L in the clearance of PtdIns4P, presumably

by transfer to and hydrolysis at the ER. If correct, this hypothesis predicts that excessive

recruitment of ORP1L would cause faster and more homogeneous clearance of PtdIns4P from

the phagolysosome. Accordingly, the rate of PtdIns4P disappearance was considerably greater

in cells expressing mCherry-tagged ORP1L than in cells transfected with unconjugated mCherry

(Fig. 5.7 b-d).

To more directly ascertain its role in PtdIns4P catabolism, we inactivated the Orp1l gene in

RAW macrophages by targeted deletion using CRISPR/Cas9. The effectiveness of the deletion

was validated at both the genomic level (Fig. 5.9a) and by immunoblotting (Fig. 5.9b). We then

proceeded to compare the kinetics of PtdIns4P clearance from phagosomes formed by wild type

and ORP1L-deficient cells (Fig. 5.9 c-f). In contrast to the wild-type cells, which displayed the

normal disappearance of the phosphoinositide at late stages of phagolysosomal maturation, the

ORP1L knockout cells cleared PtdIns4P more slowly. Despite some heterogeneity among the

knockout cells—suggestive of the existence of an alternative PtdIns4P clearance mechanism

present in varying amounts—the overall difference in the rate of disappearance was highly

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Figure 5.7. ORP1L expression accelerates late PtdIns4P disappearance in maturing phagosomes. a) Representative time-lapse confocal micrographs of RAW macrophages co-expressing mCh-2xP4M

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and GFP-ORP1L 20 min. after phagocytosis; first column of insets show magnifications of phagosomes in the main micrograph; second column of insets shows surface intensity plots corresponding to these phagosomes. b) Representative confocal micrographs of RAW macrophages co-expressing GFP-2xP4M and a control mCh vector at different times during phagosome maturation; insets show magnifications of red boxes; green frame for the GFP channel and red frame for the mCh channel c) Representative confocal micrographs of RAW macrophages co-expressing GFP-2xP4M and mCh-ORP1L at different times during phagosome maturation. d) Black dotted line: PtdIns4P dynamics during phagocytosis with expression of a control mCh vector; red line: PtdIns4P dynamics during phagocytosis with the expression of mCh-ORP1L; shown are the means of three experimental replicates of fixed cells, error bars represent SEM; ****p<0.0001 of a two-way ANOVA comparing cell lines. Scale bars = 10 µm throughout the figure.

significant (Fig. 5.9 e-f). Similar results were obtained using two other ORP1L knockout clones

(Fig. 5.10).

Figure 5.8. PtdIns4P and ORP1L localize to mutually exclusive phagosomal microdomains. a) Representative time-lapse gallery of confocal micrographs showing mutual exclusion between PtdIns4P (detected with mCh-2xP4M) and GFP-ORP1L. Scale bars = 10 µm.

Based on its domain structure, and by analogy with other proteins of the same family (de Saint-

Jean et al., 2011; Maeda et al., 2013; Tong et al., 2013; Zhao and Ridgway, 2017), we assumed

that ORP1L functions by transferring PtdIns4P to the ER and that this requires the interaction

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Figure 5.9. ORP1L KO impairs late PtdIns4P disappearance in maturing phagosomes. a) TIDE analysis showing nucleotide deletions or insertions of a homozygous ORP1L CRISPR KO clone RAW macrophage cell line. b) Representative western blot showing the expression of ORP1L in WT, control

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(cells stably expressing CAS9 and transduced with control sgRNA) and ORP1L KO RAW macrophages. c) Representative confocal micrographs of control RAW macrophages expressing GFP-2xP4M at different times during phagosome maturation. d) Representative confocal micrographs of ORP1L KO RAW macrophages expressing GFP-2xP4M at different times during phagosome maturation. e) PtdIns4P dynamics in maturing phagosomes of control cells (black bars) and ORP1L KO cells (white bars); shown are means of >140 phagosomes per time-point and per condition, acquired in three experimental replicates with fixed cells, error bars represent SEM; ****p<0.0001 of a two-way ANOVA comparing cell lines. f) Scattered plot showing normalized phagosomal GFP-2xP4M fluorescence intensity for individual phagosomes after 60 min. of phagocytosis; white circles represent control cells (184 phagosomes), gray squares represent ORP1L KO cells (201 phagosomes); horizontal lines show the mean of the total number of phagosomes of each group, measured in three independent experimental replicates, error bars represent SEM; ****p<0.0001 of an unpaired t-test comparing cell lines. Scale bars = 10 µm throughout the figure.

between its FFAT domain with VapA/B. Accordingly, in primary macrophages VapA and VAPB

were found to line the phagolysosomal membrane (Fig. 5.11b and 5.12a), where they co-

localized with GFP-ORP1L when expressed in RAW cells (Fig. 5.11c and 5.12c). This close

association was mediated by ORP1L, since it was absent when a mutant form of ORP1L lacking

the FFAT domain (designated FFAT* in Fig. 5.11) was expressed instead (Figs. 5.11c and d,

and 5.12c).

Previous studies have shown that ORP1L can sense, bind and transfer sterols (Rocha et al.,

2009; Vihervaara et al., 2011; van der Kant et al., 2013a; Wijdeven et al., 2016; Zhao and

Ridgway, 2017), which are thought to bind to the hydrophobic pocket of the ORD domain.

Interestingly, this pocket contains two conserved histidine residues that could coordinate the

binding of PtdIns4P (de Saint-Jean et al., 2011; Maeda et al., 2013; Tong et al., 2013). To test

whether the ORD domain is involved in PtdIns4P binding and transfer, we mutated this double-

histidine motif (Fig. 5.11a) and assessed the ability of the resulting mutant ORP1L (called ORP*

in Fig. 5.11) to accelerate the clearance of the phosphoinositide from phagolysosomes as was

shown for the wild-type ORP1L (Fig. 5.7 b-d). Note that in this and subsequent experiments

where mutant forms of ORP1L were expressed, the endogenous wild type ORP1L was silenced

using siRNA (Fig. 5.12b) to eliminate its countervailing effects. As illustrated in Fig. 5.11 e-f,

elimination of the double-histidine motif did not interfere with the recruitment of ORP1L-ORD* to

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Figure 5.10. Additional ORP1L KO clones impair PtdIns4P disappearance. a) TIDE analysis of a homozygous ORP1L CRISPR KO clone RAW macrophage cell line. b) Representative western blot

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showing the expression of ORP1L in WT, control (cells stably expressing CAS9 and transduced with control sgRNA) and ORP1L KO RAW macrophages. c) Representative confocal micrographs of control RAW macrophages expressing GFP-2xP4M at different times during phagosome maturation. d) Representative confocal micrographs of ORP1L KO RAW macrophages expressing GFP-2xP4M at different times during phagosome maturation.

the phagolysosomes, but eliminated its ability to stimulate the clearance of PtdIns4P, assessed

by measuring the kinetics of release of the 2xP4M biosensor.

Using the same assay, we assessed whether the FFAT domain is also required for PtdIns4P

depletion. As shown in Fig. 5.11 e-f, the FFAT* mutant was similarly incapable of accelerating

PtdIns4P disappearance, presumably because linkage with the ER was impaired.

5.3.5 ORP1L-dependent phagolysosome-to-ER PtdIns4P transport

The preceding results imply that ORP1L is involved in PtdIns4P homeostasis and suggest that

this phosphoinositide plays an important function in phagolysosome maturation. Indeed, as

documented in Fig. 5.3c, PtdIns4P-positive tubular structures emanate from the phagosomes

during the late, resolution stages. We therefore wondered whether PtdIns4P might serve to

recruit or activate molecular motors required for membrane tubulation and scission. Whereas

ORP1L forms a complex with Rab7 and RILP that recruits dynein-dynactin motors, promoting

transport towards the minus-end of microtubules (Johansson et al., 2007), the majority of the

tubules we observed using lattice light-sheet microscopy extend centrifugally, towards the

plasma membrane. We therefore hypothesized that PtdIns4P-rich structures recruit or activate

motors such as kinesins that are directed towards the plus-end of microtubules. Kinesin motors

link to lysosomes via the small GTPase Arl8B and its effector SKIP (PLEKHM2) (Rosa-Ferreira

and Munro, 2011). In this context, Arl8B has been recently implicated in the clearance of cell

corpses by C. elegans (Fazeli et al., 2018). We therefore considered the possibility that

PtdIns4P plays a role in their recruitment. Co-expression experiments showed the PtdIns4P

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Figure 5.11. ORP1L binds and transports PtdIns4P to the ER. a) Schematic representation of: WT ORP1L (top); ORP1L with a mutated FFAT motif (D478A) incapable of binding VapA and VapB in the ER

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(middle); and ORP1L with a mutated lipid-binding pocket within the ORD (HH651/652AA) predicted to abolish PtdIns4P binding. b) Localization of endogenous VapA in primary human macrophages after 40 min. of phagocytosis of HRBC; inset shows a magnification of the phagosome in the red box. c) Representative confocal micrograph of RAW macrophages co-expressing GFP-ORP1L and mCh-VapA after 30 min. of phagocytosis; insets show magnifications of extended focus of the phagosome from the main micrograph. d) Representative confocal micrograph of RAW macrophages co-expressing GFP-ORP1L-FFAT* and mCh-VapA after 30 min. of phagocytosis; insets show magnifications of extended focus of the phagosome from the main micrograph. e) Representative confocal micrographs after 30 min of phagocytosis of: control RAW macrophages co-transfected with GFP-2xP4M and an mCh-vector and treated with scrambled siRNA (first column); in the following columns RAW macrophages were treated with siRNA targeting ORP1L and co-transfected with FP-2xP4M and, either a control vector or the different ORP1L constructs tagged with fluorescent proteins. f) The graph shows normalized FP-2xP4M phagosomal intensity of individual phagosomes of RAW macrophages under the conditions summarized in e). Scale bars = 10 µm throughout the figure.

sensor 2xP4M to co-localize with Arl8B or SKIP on the phagosomal membrane (Fig. 5.13 a-b

and 5.14 a-b) and in the tubular structures emanating therefrom. In sharp contrast, mutual

exclusion was observed when comparing the distribution of ORP1L with either Arl8B or SKIP

(5.13d and e). The pattern of exclusion was highly reminiscent of that observed for ORP1L vs.

PtdIns4P (Fig. 5.7a).

While Arl8B does not contain a lipid-binding domain, SKIP has been reported to contain a PH

domain near its C-terminus (Boucrot et al., 2005). PH domains are noted for their ability to bind

phosphoinositides with high affinity (Lemmon, 2003; Hammond and Balla, 2015). Because of

this, we hypothesized that PtdIns4P might recruit SKIP to the phagolysosomal membrane. To

address this, first we performed a lipid overlay assay with purified recombinant GST-tagged

SKIP on a membrane spotted with decreasing concentrations of phosphoinositides. Notably, at

low concentrations, GST-SKIP bound mono-phosphorylated phosphoinositides with higher

affinity. High SKIP concentrations also bound to bisphosphorylated PIPs (Fig. 5.13c and 5.14c).

To test the functional implications of this interaction we compared the accumulation of SKIP in

phagosomes under conditions where PtdIns4P levels were altered. To lower the phagosomal

PtdIns4P we overexpressed wild type ORP1L. Conversely, we used the ORP1L-deleted cells to

increase the levels of the phosphoinositide. Macrophages were challenged with IgG-SRBC and

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Figure 5.12. ORP1L – VAPA/B interactions mediate phagosome – ER contacts. a) Localization of endogenous VapB in primary human macrophages after 40 min of phagocytosis of IgG-HRBC. b) Relative mRNA levels measured by qPCR of control RAW cells treated with scrambled siRNA cells (black bar) and cells treated with siRNA against Orp1l; means + SD of six independent experiments each one normalized to mRNA levels of cells treated with scrambled siRNA. c – h. Maximum intensity projections of RAW cells after 30 min of phagocytosis of SRBC expressing: c) GFP-ORP1L and mCh-VapA; d) GFP-ORP1L-FFAT* and mCh-VapA; e) mCh-ORP1L-ORD* and GFP-VapA; f) GFP-ORP1L and mCh-VapB; g) GFP-ORP1L-FFAT* and mCh-VapB; and h) GFP-ORP1L-ORD* and mCh-VapB. Insets show magnifications of phagosomes in each channel and their merge.

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Figure 5.13. PtdIns4P directly binds SKIP and recruits it to the phagolysosome. a) Representative confocal micrograph of RAW macrophages co-expressing GFP-2xP4M and mCh-Arl8b after 40 min. of phagocytosis; insets show magnifications of the phagosome within the white box; the graph below represents a line-scan of fluorescent intensities along the periphery of the phagosome. b) Representative confocal micrograph of RAW macrophages co-expressing mCh-2xP4M and GFP-SKIP after 40 min. of

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phagocytosis; insets show magnifications of the phagosome within the white box; the graph below represents a line-scan of fluorescent intensities along the periphery of the phagosome. c) Protein – lipid overlay assay; a hydrophobic membrane spotted with the indicated lipids and overlaid with purified indicated proteins; bound protein was detected by immune-blotting. d) Representative confocal micrograph of RAW macrophages co-expressing GFP-ORP1L and mCh-Arl8b after 40 min. of phagocytosis; insets show magnifications of the phagosome within the white box; the graph below represents a line-scan of fluorescent intensities along the periphery of the phagosome. e) Representative confocal micrograph of RAW macrophages co-expressing mCh-ORP1L and GFP-SKIP after 40 min. of phagocytosis; insets show magnifications of the phagosome within the white box; the graph below represents a line-scan of fluorescent intensities along the periphery of the phagosome. f) RAW macrophages expressing GFP-SKIP after 45 min of phagocytosis of IgG-SRBC under: control conditions (left panel); overexpression of mCh-ORP1L (middle panel); and ORP1L KO (right panel). g) Quantitation of SKIP recruitment to phagosomes under the conditions shown in f. Shown are means + SEM of mean fluorescent intensities of GFP-SKIP in phagosomal membranes.

phagosomes allowed to mature for 45 min. Phagosomes that accumulated overexpressed

ORP1L (and hence likely depleted of PtdIns4P) recruited lower amounts of SKIP (Fig. 5.13f and

g). Conversely, phagosomes lacking ORP1L acquired significantly more SKIP than those of

control cells (Fig. 5.13f and g). We used a similar strategy to assess the dependence of Arl8B

on phagosomal PtdIns4P. Arl8B levels also decreased in phagosomes with excess ORP1L,

although no significant difference was noted in the ORP1L knockout cells (Fig. 5.14 and e).

Our data suggested a link between PtdIns4P and the tubules and vesicles that seemingly

mediate phagosome resolution. To evaluate this possibility, we analyzed the effects of altering

PtdIns4P on phagolysosome remodeling. Phagosome size and integrity were assessed as in

Fig. 1. As before, untreated macrophages showed extensive phagolysosome fragmentation 4.5

hours after ingestion (Fig. 5.15 a and b). When PtdIns4P was depleted by overexpressing

ORP1L, the phagosomes remained considerably larger, in some instances exceeding the

original phagosomal size (Fig. 5.15a and b). Of note, the contents of these phagosomes were

clearly degraded, implying that fusion with lysosomes did occur. Interestingly, in macrophages

of ORP1L-deficient macrophages ≈50% of the phagosomes appeared to have resolved, while

the remaining half had a size that was similar to or larger than the nascent phagosomes (Fig.

5.15a and b). A role for PtdIns4P during resolution was further supported by lattice light-sheet

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Figure 5.14. PtdIns4P stabilizes the Arl8b – SKIP interaction in the phagosomal membrane. a) Representative confocal micrographs of phagosomes in RAW macrophages transfected with a) GFP-SKIP and mTq2-2xP4M; or b) mCh-Arl8b and GFP-2xP4M. c) Protein – lipid overlay assay; a hydrophobic membrane spotted with the indicated lipids and overlaid with purified recombinant GST-SKIP; bound protein was detected by immune-blotting. d) RAW macrophages expressing mCh-Arl8b after 45 min of phagocytosis of IgG-SRBC under: control conditions (left panel); overexpression of mCh-ORP1L (middle panel); and ORP1L KO (right panel). e) Quantitation of Arl8b recruitment to phagosomes under the conditions shown in f. Shown are means + SEM of mean fluorescent intensities of mCh-Arl8b in phagosomal membranes.

microscopy observations. As shown in Fig. 5.15c and, the volume of phagosomes was reduced

as PtdIns4P-positive tubules extended.

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Figure 5.15. ORP1L mediates phagosome resolution by regulating phagosomal PtdIns4P. a) RAW macrophages were challenged with TAMRA-SRBC and phagosomes allowed to mature for 4.5 h: macrophages were transfected with a control GFP vector (top); with WT GFP-ORP1L (middle); and ORP1L KO macrophages were transfected with a GFP vector (bottom); scale bars = 10 µm. b) The graph shows the major diameters of individual vacuoles containing TAMRA signal after 4.5 of phagocytosis in RAW macrophages with different ORP1L levels; the gray region limited by dotted red lines shows the size range of intact SRBC-containing phagosomes. c) Time-lapse lattice light-sheet micrographs of RAW macrophages expressing GFP-2xP4M before, during and after tubulation events associated with resolution. d) Time-lapse lattice light-sheet micrographs of RAW macrophages expressing GFP-ORP1L at the same time frame represented in c); scale bars = 3 µm.

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5.4 Discussion

Following degradation of their contents, phagosomes must be resorbed to recycle their

components and to vacate space for additional rounds of phagocytosis. Surprisingly, the

concluding stage of phagocytosis remains largely unexplored and poorly defined. An mTOR-

dependent step has been described, where phagosomal fission events redistribute membrane

and contents into the lysosomal network (Krajcovic et al., 2013). In addition, antigens are

delivered from phagosomes to the membrane for presentation, a process similarly linked to

tubule formation and generation of transport vesicles (Mantegazza et al., 2014; Saric et al.,

2016). The molecular basis of the tubulation and fragmentation processes, however, are not

well defined.

More mechanistic information has been gained for the analogous process of autolysosome

remodeling. Like the phagolysosomes of myeloid cells, autophagosomes emit tubules that are

involved in the reformation of lysosomes. Clathrin and PtdIns(4,5)P2 are central to this process

(Rong et al., 2012). Specifically, PtdIns(4,5)P2 was proposed to directly interact with the kinesin

motor KIF5A to drive tubulation (Du et al., 2016). In macrophages, however, PtdIns(4,5)P2 is

not detectable in resolving phagolysosomes (Fig 5.16), where PtdIns4P is instead the

predominant phosphoinositide. Indeed, at its peak, the concentration of PtdIns4P in the

phagolysosome greatly exceeds that in the plasma membrane (Fig. 5.3b). Interestingly, a

recent study established the importance of PtdIns5P during autolysosome remodeling. Our

observation that SKIP can also bind PtdIns5P (Fig. 5.13c) may have relevance in this context.

Although there are significant caveats associated with these type of lipid binding assays.

Following its abrupt disappearance shortly after phagosome closure, PtdIns4P reappears

concomitantly with the elimination of PtdIns3P marking the early to late phagosome transition

[see (Levin et al., 2017) and Fig. 5.3b]. Reacquisition of PtdIns4P is mediated by PI4K2A—the

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Figure 5.16. PtdIns(4,5)P2 is not present in phagosomes during PtdIns4P-positive tubulation. a) Representative confocal micrographs of RAW macrophages expressing GFP-2xP4M after 45 min of phagocytosis and stained with phalloidin conjugated to Alexa Fluor 568. Each row shows different cells under the same conditions. b) Representative confocal micrographs of RAW macrophages co-expressing GFP-WASH and mCh-2xP4M after 45 min of phagocytosis. c) Representative confocal micrographs of RAW macrophages expressing GFP-PH-PLCδ after 45 min of phagocytosis and stained with phalloidin conjugated to Alexa Fluor 568. Insets throughout the figure are magnifications of individual phagosomes. Scale bars = 10 µm.

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kinase responsible for its synthesis from PtdIns (Jeschke et al., 2015; Levin et al., 2017). Here

we report an additional, late phase of PtdIns4P depletion that coincides with the onset of

phagosome resolution. We believe that at least two components contribute to the late PtdIns4P

disappearance stage: ORP1L recruited by Rab7 transfers a fraction of the PtdIns4P to the ER,

where it is hydrolyzed by SAC1. The distribution of ORP1L on the phagosomal membrane is

not continuous, forming patches where PtdIns4P is preferentially depleted. The remainder of

the membrane, which retains PtdIns4P, is the source of the tubules that mediate the resolution

process. Detachment of such PtdIns4P-enriched tubules contributes the second component to

the late phagolysosomal PtdIns4P depletion event. The existence of at least two separate

components explains why a fraction of the PtdIns4P clearance persists in the ORP1L knockout

cells.

The finding that ORP1L transfers PtdIns4P to the ER is somewhat unexpected, inasmuch as

ORP1L has been widely documented to be a cholesterol sensor and transporter. However, the

reported ability of its ORD domain to bind PtdIns4P with even greater affinity than sterols is

consistent with our findings. Whether the transport step is unidirectional is not presently clear.

OSBP and other related proteins mediate counter-transport mechanisms, and it is conceivable

that another lipid is delivered from the ER to the phagolysosome in exchange for PtdIns4P.

One obvious candidate would be cholesterol. In fact, a late phase of reacquisition of cholesterol

by phagosomes has been reported (Rai et al., 2016). We attempted to demonstrate a role for

ORP1L in this process, but failed to reliably detect dynamic changes in cholesterol in late

phagolysosomes. Nevertheless, the existence of a counter-transport mechanism, dependent on

the steep PtdIns4P chemical gradient, remains a viable possibility that requires additional study.

While several studies have reported essential roles of PtdIns4P at the Golgi and plasma

membrane, little is known about its function in the endolysosomal network. In the recycling

pathway PtdIns4P regulates retromer function and has been linked to actin nucleation via

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WASH (Niu et al., 2013; Dong et al., 2016; Marquer et al., 2016). The latter is necessary for

scission of retromer-induced tubules from endosomes. Accordingly, the PtdIns4P-rich regions

of phagolysosomal membranes that we observed were both actin- and WASH-positive (not

shown). However, our evidence suggests that PtdIns4P plays a role not only in the scission of

tubules, but also in their formation. We propose a mechanism whereby by binding SKIP,

PtdIns4P stabilizes the Arl8B-SKIP complex on the phagolysosomal surface, enabling tubule

extension and transport towards the cell periphery.

Our model relies on the exquisite temporal and spatial segregation of PtdIns4P-enriched

regions within the phagosomal membrane. Earlier elegant studies have in fact shown the

existence of phagosomal microdomains: dynein and Rab7 were found to cluster in cholesterol-

rich patches in phagolysosomes (Rai et al., 2016). These domains are presumably also

enriched in the Rab7 effectors ORP1L and RILP. Our observation that ORP1L depletes

PtdIns4P from these microdomains while adjacent regions retain the phosphoinositide raises

the possibility that opposing forces are required for tubule extrusion. Specifically, while

PtdIns4P-enriched domains experience centrifugal force exerted by the Arl8B/SKIP/kinesin

complex, the Rab7-containing patches may be driven in the opposite direction, i.e. centripetally,

by RILP/dynein (Rai et al., 2016), depending on the prevailing concentration of cholesterol

(Vihervaara et al., 2011). At the Rab7-rich domains ORP1L not only depletes PtdIns4P, but

forms contact sites with the ER that provide physical tethering to enable the extension and

scission of tubules by microtubule-associated motors, without dragging the entire organelle to

the cell periphery when centrifugal motors are dominant.

In summary, we showed a new phase of PtdIns4P metabolism during the resolution stage of

phagolysosomes and were able to attribute at least a fraction of the change to transfer of the

inositide to the ER via ORP1L. In addition, we documented the formation of defined PtdIns4P

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microdomains that, by associating with SKIP/Arl8B, promote the generation of tubules that

contribute to the resolution of phagolysosomes.

Figure 5.17. Working model. Middle panel) phagolysosomes have an uneven PtdIns4P distribution in their membranes; tubular and vesicular structures emanate from PtdIns4-rich areas, while adjacent areas are in contact with the ER. Right panel) PtdIns4P is depleted in phagolysosome – ER contact sites. ORP1L transfers PtdIns4P to the ER, where the lipid is degraded by SAC1. Left panel) In areas of the phagolysosome where it is enriched, PtdIns4P stabilizes the Arl8b-SKIP-Kinesin complex by binding and recruiting SKIP to the phagosomal membrane. This promotes the generation of the aforementioned tubular and vesicular structures.

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Chapter 6 General Discussion

Summary of findings, future directions and concluding 6remarks

The work described in this dissertation primarily aimed to elucidate potential roles of

phosphoinositides during late stages of phagocytosis, namely late phagosome maturation and

phagolysosome resolution. Specifically, this work advances the understanding of

phosphoinositides during phagocytosis by addressing the following questions: 1) What are the

dynamics of PtdIns4P during each stage of phagocytosis? 2) What are the enzymes and

proteins responsible for PtdIns4P metabolism during phagocytosis? 3) What are the functional

implications of the presence of PtdIns4P in the phagosomal membrane? And 4) How does the

phosphoinositide exert these functions?

6.1 PtdIns4P dynamics during phagocytosis

6.1.1 Summary of findings

In Chapter 4, I describe multiphasic dynamics of PtdIns4P levels in phagosomes during

formation and maturation. Specifically, with the use of a PtdIns4P-sensing probe, I determined

the localization of this phosphoinositide during phagocytosis with temporal, suborganellar

resolution. I showed that as phagosomes form, PtdIns4P levels transiently increase in the

phagocytic cup, followed by an abrupt decline as the maturation of the compartment initiates.

This disappearance coincides with the acquisition of Rab5 and production of PtdIns3P in the

early phagosome. Phagosomes lose both Rab5 and PtdIns3P during the transition from early

to late maturation. The late stage is characterized by the acquisition of active Rab7; it is during

this period that PtdIns4P gradually accumulates in the phagosomal membrane. Notably,

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PtdIns4P detection in late phagosomes / phagolysosomes is noticeably higher than in the PM.

In Chapter 5, I identified yet another metabolic stage of PtdIns4P during phagocytosis. This

change occurs during the transition from the biogenesis of phagolysosomes to the initiation of

their resolution and it consists of a gradual disappearance of the phosphoinositide from the

phagosomal membrane. Intriguingly, this disappearance is accompanied by the generation of

microdomains where PtdIns4P was enriched in some areas of the phagosomal membrane

while becoming partially or completely depleted in others.

6.1.2 Future directions

Several open questions were generated by these observations. Among them, distribution and

relative abundance of other phosphoinositides, especially PtdIns(3,5)P2, in phagosomes during

the phagolysosomal stage is of particular interest. PtdIns(3,5)P2 is thought to be functional the

lysosomal level but thorough characterizations have not yet been performed due to the lack of

reliable sensing probes. Additionally, my studies suggest that the presence of PtdIns3P and

PtdIns4P in phagosomes is mutually exclusive; it is tantalizing to analyze whether the

metabolism of these two lipids is coupled and coordinated. Further studies can also address

unequivocally whether the transition from the PtdIns3P- to the PtdIns4P-positive stage is a

cause or a consequence of phagosome maturation. Finally, since PtdIns4P synthesis in the

late phagosome coincides with the activation of Rab7 it will be important to address the

potential interaction between them. This will be discussed in subsection 6.3.1.

6.2 PtdIns4P metabolism mechanisms during phagocytosis

6.2.1 Summary of findings and future directions

Initial transient PtdIns4P increase in the forming phagosome

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The plasmalemmal pool of PtdIns4P is generated by the activity of PI4KA. This cytosolic lipid

kinase transiently localizes to the PM in a complex with TTC7, FAM126 and EFR3.

Consistently, we found the kinase localizing to forming phagocytic cups, suggesting this may be

the source of the PtdIns4P enrichment in the extending pseudopods. It is known that local

increases in PtdIns(4,5)P2 levels are necessary for phagocytosis. PtdIns4P is arguably the

substrate for this increase; hence the PtdIns4P increase I described is presumably necessary

to sustain these changes in lipid levels. It becomes compelling to investigate whether the

recruitment and activity of PI4KA are essential for phagocytosis. Moreover, how is the

recruitment of the complex regulated in the phagocytic cup? Additionally, the question remains

whether PtdIns4P plays any additional roles beyond that of a precursor at this level.

Previous studies suggest that some PtdIns4P is generated from the degradation of

PtdIns(4,5)P2 by the 5-phosphatases OCRL and INPP5B, which are recruited to the nascent

phagosome in Rab5-positive vesicles. It is conceivable then, that the ‘spike’ of PtdIns4P

observed during scission of the phagosome is the result of the action of these 5-phosphatases.

Sharp PtdIns4P disappearance during the early maturation

Perhaps one of the most striking observations in this work was the drastic disappearance of

PtdIns4P—within 5 to 15 s—as phagosomes were internalized. Interrogating the mechanism

behind this stage was, however, a challenging endeavor. While we could not elucidate all the

enzymes involved in this multi-factorial phenomenon, I was able to identify two key players in

the catabolism of PtdIns4P: SAC2, a then recently characterized 4-phosphatase; and PLC

enzymes, primarily known for hydrolyzing PtdIns(4,5)P2. Adding to the complexity of this

process may be the dynamic remodeling of the phagosomal membrane immediately after

internalization. Notably, in this regard, while the early phagosome is presumably enriched in

PtdIns, to the best of my knowledge there are no PtdIns4P-synthesizing enzymes in this

compartment. Because SAC2 is delivered in Rab5 compartments, it may be coordinated with

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the acquisition of Vps34, accounting for the mutual exclusion of PtdIns4P and PtdIns3P at this

stage. It is thus of interest to understand whether prevention of PtdIns4P disappearance

arrests maturation and the generation of PtdIns3P, or whether both phosphoinositides can co-

exist in the same organelle.

Gradual PtdIns4P reacquisition during the late maturation

Functionally, the generation of PtdIns4P in the late phagosome is arguably the most

functionally important phase of its dynamics. My work showed that PI4K2A—a lipid kinase

localized in the TGN and endolysosomal system—is responsible for the synthesis of the vast

majority of PtdIns4P in maturing phagosomes. This kinase is delivered to the endolysosomal

system from the TGN in an AP-3 inter-dependent manner. This raises the possibility that it is

involved in the delivery of degradative enzymes to the phagosome. Additionally, PI4K2A

deficiency impaired phagosomal acidification. As mentioned earlier, one of the most relevant

open questions lays in a potential relationship between the acquisition of Rab7 and PtdIns4P by

phagosomes and their potential joint roles through the late maturation stages. Because my

work suggests that PI4K2A remains in the maturing phagosome for at least an hour, it becomes

relevant to analyze how its activity is regulated and how this relates to the late disappearance

stage of PtdIns4P.

Late PtdIns4P disappearance from the phagolysosome

In Chapter 5, I describe a late PtdIns4P disappearance phase. As mentioned before, this stage

resulted in the generation of membrane microdomains. However, this raises a fundamental

question: how is a lipid, which is typically predicted to diffuse freely throughout a membrane,

confined and enriched in specific domains while depleted from others?

My work identified ORP1L —a putative Rab7 effector— as a PtdIns4P-transport protein. I

provide evidence that the direction of the lipid transfer is from the phagolysosome towards the

ER where I hypothesize that the phosphoinositide is rapidly degraded by SAC1. Tellingly,

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PtdIns4P transfer was dependent on the capacity of ORP1L to tether the phagosome to the ER

through the interaction of an FFAT motif with the ER-resident proteins VAPA and/or VAPB.

Additionally, two conserved histidine residues in the ORD of ORP1L coordinated binding of

PtdIns4P. Therefore, the model I propose supports SAC1 action in cis (in the ER membrane)

rather than in trans (‘reaching’ across and acting on another membrane) as it has been

proposed in models of other biological processes. Although working with SAC1 has significant

limitations, future experiments should aim to determine unequivocally its role in my hypothesis.

The observation that the disappearance of PtdIns4P was only partially impaired in ORP1L KO

macrophages suggests that an additional PtdIns4P degradation mechanism exists at this stage.

This notion is supported by the data that showed no increase in the maximum levels of

phagosomal PtdIns4P in these cells. Because of this, it seems relevant to explore additional

mechanisms that regulate PtdIns4P levels at late stages.

Another intriguing future direction of my work relates to a potential counter-transport

mechanism whereby another lipid species can be transported from the ER to the

phagolysosome. Indeed, several studies suggest that PtdIns4P gradients between organelle

membranes and the ER act as the driving force for counter-transport of sterols and other lipids

such as phosphatidylserine. Because ORP1L can bind and transport cholesterol, it is possible

that under certain conditions the sterol can be exchanged for PtdIns4P. Although we were not

successful in the detection of cholesterol with enough sensitivity to discern microdomains of the

phagosomal membrane, future studies can further explore this idea.

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6.3 Functional implications of PtdIns4P during phagocytosis

6.3.1 Summary of findings and future directions

An important objective of this dissertation was the definition of potential roles of PtdIns4P

during phagocytosis. In both Chapters 4 and 5, I provide different lines of evidence for

functions of the phosphoinositide during late stages of phagocytosis. First, partial depletion of

PI4K2A and significant decreases in PtdIns4P levels in maturing phagosomes impaired

acidification of the compartment—a hallmark of phagosome maturation. However, it is not clear

whether this was due to lack of lysosomal fusion with phagosomes or whether it was because

PtdIns4P directly or indirectly regulates the acidification of phagolysosomes. The first

possibility is supported by studies in autophagosomes where lack of PtdIns4P prevents

lysosome–autophagosome fusion. Evidence potentially supporting the latter option is based on

interactions of a V-ATPase subunit with PtdIns4P in yeast. Future experiments should aim to

define how depletion of PtdIns4P impairs phagosome acidification. This may have implications

as to how some bacterial species such as Coxiella burnetti and Legionella pneumophila evade

elimination by certain phagocytes.

The second PtdIns4P function that my work revealed, deals with the activation state of Rab7 in

the phagosomal membrane and / or its ability to recruit effector proteins. Indeed, acute

depletion of PtdIns4P from the phagosomal membrane through the controlled recruitment of a

chimeric phosphatase prevented the acquisition of RILP, a putative active-Rab7 effector. While

these data suggest that PtdIns4P is important for the activation of the GTPase, their

relationship may be more complex. In early compartments, active Rab5 recruits Vps34 that in

turn produces PtdIns3P. The coordinated action of Rab5 and PtdIns3P recruit specific effectors

and controls the maturation of these compartments. It is conceivable that Rab7 may be

involved in the production of PtdIns4P in the phagosomal membrane, and by analogy with the

example above, they may coordinate the recruitment of effectors that direct the late maturation.

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In Chapter 5, I provide evidence of a role for PtdIns4P during tubulation of phagolysosomes.

This is likely through the interaction I described between the phosphoinositide and the kinesin-

binding protein SKIP. Hence, my model proposes that PtdIns4P present in microdomains of

the phagolysosome aids Arl8B in the recruitment of SKIP. In order to generate tubules, this

complex links these regions of the membrane to kinesin motors that interact with microtubules.

Further supporting this model is the evidence that I provide showing that dysregulation of

PtdIns4P levels in phagolysosomes impaired the resolution of the compartments. This may be

a result of the phagosome’s inability to tubulate in order to initiate the fission events that

characterize resolution.

It does remain a question whether this mechanism is related to antigen presentation. Yet,

integrating diverse lines of evidence makes PtdIns4P a suitable candidate for involvement in

this process. As mentioned earlier, PI4K2A is delivered to endosomes and lysosomes in an

AP-3-dependent manner. Antigen presentation is impaired in dendritic cells of AP-3 KO mice.

Moreover, Arl8B has also been described as necessary for antigen presentation. Incorporating

these sets of observations led me to the rationale that delivery of PI4K2A in AP-3 vesicles is

required for production of PtdIns4P in phagosomes. In turn, the phosphoinositide coordinates

the recruitment of machinery necessary for antigen presentation (e.g. Arl8B). This hypothesis

is yet to be tested. Based on our observations one would predict that affecting any step

downstream of PtdIns4P production (e.g. AP-3) or components that are directly or indirectly

regulated by the lipid (i.e. SKIP, Arl8B and kinesin motors) in the phagolysosome, would

severely impair phagosome resolution.

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6.4 Concluding remarks

Phagocytosis is a critical process in immunity and homeostasis. It represents a complex set of

events involving a vast compendium of molecules that initiates with the surveillance of the

extracellular environment and concludes with the reformation of lysosomes—the so-called

‘terminal’ organelles of cells. Due to this ‘evolving’ nature, phagocytosis is subject to a tight—

yet exquisite—regulation in space and time. Substantial literature has established paramount

roles for phosphoinositides as crucial orchestrators of the endocytic process. Indeed,

manipulations of their metabolism significantly impair different aspects of phagocytosis.

Diverse roles have been established for these versatile phospholipids during the process,

mainly through the recruitment of effector proteins. While these phenomena have been

extensively characterized for phagosome formation and the early maturation, our understanding

of signaling regulation during the late phagosome maturation and resolution stages is still

limited. My work defined the metabolic changes and mechanisms of PtdIns4P action during

each stage of phagocytosis; additionally, I provided evidence that suggests diverse roles of this

phosphoinositide during late stages of the process (i.e., late maturation, phagolysosome

biogenesis and resolution). My work also highlights the importance of studying and

understanding the resolution of phagosomes—a crucial, yet underappreciated, stage of

phagocytosis—in order to both resume the immune response and conclude the ‘waste

management’ process for homeostatic purposes in an efficient manner.

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References Abella, J. V. G., Galloni, C., Pernier, J., Barry, D. J., Kjær, S., Carlier, M.-F., and Way, M. (2016). Isoform diversity in the Arp2/3 complex determines actin filament dynamics. Nat. Cell Biol. 18, 76–86.

Abou-Kheir, W., Isaac, B., Yamaguchi, H., and Cox, D. (2008). Membrane targeting of WAVE2 is not sufficient for WAVE2-dependent actin polymerization: a role for IRSp53 in mediating the interaction between Rac and WAVE2. J. Cell Sci. 121, 379–390.

Abraham, R. T. (2015). Cell biology. Making sense of amino acid sensing. Science 347, 128–129.

Alloatti, A. et al. (2015). Toll-like Receptor 4 Engagement on Dendritic Cells Restrains Phago-Lysosome Fusion and Promotes Cross-Presentation of Antigens. Immunity 43, 1087–1100.

Amann, K. J., and Pollard, T. D. (2001). Direct real-time observation of actin filament branching mediated by Arp2/3 complex using total internal reflection fluorescence microscopy. Proc. Natl. Acad. Sci. U. S. A. 98, 15009–15013.

Ambruso, D. R. et al. (2000). Human neutrophil immunodeficiency syndrome is associated with an inhibitory Rac2 mutation. Proc. Natl. Acad. Sci. U. S. A. 97, 4654–4659.

Anand, R. J., Kohler, J. W., Cavallo, J. A., Li, J., Dubowski, T., and Hackam, D. J. (2007). Toll-like receptor 4 plays a role in macrophage phagocytosis during peritoneal sepsis. J. Pediatr. Surg. 42, 927–932; discussion 933.

Araki, N., Hatae, T., Furukawa, A., and Swanson, J. A. (2003). Phosphoinositide-3-kinase-independent contractile activities associated with Fcgamma-receptor-mediated phagocytosis and macropinocytosis in macrophages. J. Cell Sci. 116, 247–257.

Araki, N., Johnson, M. T., and Swanson, J. A. (1996). A role for phosphoinositide 3-kinase in the completion of macropinocytosis and phagocytosis by macrophages. J. Cell Biol. 135, 1249–1260.

Arandjelovic, S., and Ravichandran, K. S. (2015). Phagocytosis of apoptotic cells in homeostasis. Nat. Immunol. 16, 907–917.

Azzoni, L., Kamoun, M., Salcedo, T. W., Kanakaraj, P., and Perussia, B. (1992). Stimulation of Fc gamma RIIIA results in phospholipase C-gamma 1 tyrosine phosphorylation and p56lck activation. J. Exp. Med. 176, 1745–1750.

Backer, J. M. (2008). The regulation and function of Class III PI3Ks: novel roles for Vps34. Biochem. J. 410, 1–17.

Bajno, L., Peng, X. R., Schreiber, A. D., Moore, H. P., Trimble, W. S., and Grinstein, S. (2000). Focal exocytosis of VAMP3-containing vesicles at sites of phagosome formation. J. Cell Biol. 149, 697–706.

Bakowski, M. A., Braun, V., Lam, G. Y., Yeung, T., Heo, W. D., Meyer, T., Finlay, B. B., Grinstein, S., and Brumell, J. H. (2010). The phosphoinositide phosphatase SopB manipulates

153

membrane surface charge and trafficking of the Salmonella-containing vacuole. Cell Host Microbe 7, 453–462.

Balce, D. R., and Yates, R. M. (2017). Fluorometric Approaches to Measuring Reductive and Oxidative Events in Phagosomes. In: Phagocytosis and Phagosomes: Methods and Protocols, ed. R. Botelho, New York, NY: Springer New York, 215–225.

Balla, A., Kim, Y. J., Varnai, P., Szentpetery, Z., Knight, Z., Shokat, K. M., and Balla, T. (2008). Maintenance of hormone-sensitive phosphoinositide pools in the plasma membrane requires phosphatidylinositol 4-kinase IIIalpha. Mol. Biol. Cell 19, 711–721.

Balla, A., Tuymetova, G., Barshishat, M., Geiszt, M., and Balla, T. (2002). Characterization of type II phosphatidylinositol 4-kinase isoforms reveals association of the enzymes with endosomal vesicular compartments. J. Biol. Chem. 277, 20041–20050.

Balla, A., Tuymetova, G., Tsiomenko, A., Várnai, P., and Balla, T. (2005). A plasma membrane pool of phosphatidylinositol 4-phosphate is generated by phosphatidylinositol 4-kinase type-III alpha: studies with the PH domains of the oxysterol binding protein and FAPP1. Mol. Biol. Cell 16, 1282–1295.

Balla, T. (2013). Phosphoinositides: tiny lipids with giant impact on cell regulation. Physiol. Rev. 93, 1019–1137.

Baskin, J. M. et al. (2016). The leukodystrophy protein FAM126A (hyccin) regulates PtdIns(4)P synthesis at the plasma membrane. Nat. Cell Biol. 18, 132–138.

Beemiller, P., Zhang, Y., Mohan, S., Levinsohn, E., Gaeta, I., Hoppe, A. D., and Swanson, J. A. (2010). A Cdc42 activation cycle coordinated by PI 3-kinase during Fc receptor-mediated phagocytosis. Mol. Biol. Cell 21, 470–480.

Bengtsson, T., Jaconi, M. E., Gustafson, M., Magnusson, K. E., Theler, J. M., Lew, D. P., and Stendahl, O. (1993). Actin dynamics in human neutrophils during adhesion and phagocytosis is controlled by changes in intracellular free calcium. Eur. J. Cell Biol. 62, 49–58.

Berger, P., Schaffitzel, C., Berger, I., Ban, N., and Suter, U. (2003). Membrane association of myotubularin-related protein 2 is mediated by a pleckstrin homology-GRAM domain and a coiled-coil dimerization module. Proc. Natl. Acad. Sci. U. S. A. 100, 12177–12182.

Beyer, M. et al. (2012). High-resolution transcriptome of human macrophages. PloS One 7, e45466.

Bianconi, E. et al. (2013). An estimation of the number of cells in the human body. Ann. Hum. Biol. 40, 463–471.

Binker, M. G. et al. (2007). Arrested maturation of Neisseria-containing phagosomes in the absence of the lysosome-associated membrane proteins, LAMP-1 and LAMP-2. Cell. Microbiol. 9, 2153–2166.

Blander, J. M., and Medzhitov, R. (2004). Regulation of phagosome maturation by signals from toll-like receptors. Science 304, 1014–1018.

Blander, J. M., and Medzhitov, R. (2006). On regulation of phagosome maturation and antigen presentation. Nat. Immunol. 7, 1029–1035.

154

Bloomfield, G., Traynor, D., Sander, S. P., Veltman, D. M., Pachebat, J. A., and Kay, R. R. (2015). Neurofibromin controls macropinocytosis and phagocytosis in Dictyostelium. ELife 4.

Bohdanowicz, M., Balkin, D. M., De Camilli, P., and Grinstein, S. (2012a). Recruitment of OCRL and Inpp5B to phagosomes by Rab5 and APPL1 depletes phosphoinositides and attenuates Akt signaling. Mol. Biol. Cell 23, 176–187.

Bohdanowicz, M., Balkin, D. M., De Camilli, P., and Grinstein, S. (2012b). Recruitment of OCRL and Inpp5B to phagosomes by Rab5 and APPL1 depletes phosphoinositides and attenuates Akt signaling. Mol. Biol. Cell 23, 176–187.

Bohdanowicz, M., Cosío, G., Backer, J. M., and Grinstein, S. (2010). Class I and class III phosphoinositide 3-kinases are required for actin polymerization that propels phagosomes. J. Cell Biol. 191, 999–1012.

Bohdanowicz, M., and Grinstein, S. (2013). Role of phospholipids in endocytosis, phagocytosis, and macropinocytosis. Physiol. Rev. 93, 69–106.

Bohdanowicz, M., Schlam, D., Hermansson, M., Rizzuti, D., Fairn, G. D., Ueyama, T., Somerharju, P., Du, G., and Grinstein, S. (2013). Phosphatidic acid is required for the constitutive ruffling and macropinocytosis of phagocytes. Mol. Biol. Cell 24, 1700–1712, S1-7.

Bonifacino, J. S., and Hurley, J. H. (2008). Retromer. Curr. Opin. Cell Biol. 20, 427–436.

Booth, J. W., Kim, M.-K., Jankowski, A., Schreiber, A. D., and Grinstein, S. (2002). Contrasting requirements for ubiquitylation during Fc receptor-mediated endocytosis and phagocytosis. EMBO J. 21, 251–258.

Borgman, K. J. E., van Zanten, T. S., Manzo, C., Cabezón, R., Cambi, A., Benítez-Ribas, D., and Garcia-Parajo, M. F. (2014). Priming by chemokines restricts lateral mobility of the adhesion receptor LFA-1 and restores adhesion to ICAM-1 nano-aggregates on human mature dendritic cells. PloS One 9, e99589.

Botelho, R. J., Teruel, M., Dierckman, R., Anderson, R., Wells, A., York, J. D., Meyer, T., and Grinstein, S. (2000). Localized biphasic changes in phosphatidylinositol-4,5-bisphosphate at sites of phagocytosis. J. Cell Biol. 151, 1353–1368.

Boucrot, E., Henry, T., Borg, J.-P., Gorvel, J.-P., and Méresse, S. (2005). The intracellular fate of Salmonella depends on the recruitment of kinesin. Science 308, 1174–1178.

Boura, E., and Nencka, R. (2015). Phosphatidylinositol 4-kinases: Function, structure, and inhibition. Exp. Cell Res. 337, 136–145.

Bray, D. (2001). Cell Movements: From Molecules to Motility, Garland Science.

Bretscher, A., Edwards, K., and Fehon, R. G. (2002). ERM proteins and merlin: integrators at the cell cortex. Nat. Rev. Mol. Cell Biol. 3, 586–599.

Bröcker, C., Kuhlee, A., Gatsogiannis, C., Balderhaar, H. J. kleine, Hönscher, C., Engelbrecht-Vandré, S., Ungermann, C., and Raunser, S. (2012). Molecular architecture of the multisubunit homotypic fusion and vacuole protein sorting (HOPS) tethering complex. Proc. Natl. Acad. Sci. U. S. A. 109, 1991–1996.

155

Brombacher, E., Urwyler, S., Ragaz, C., Weber, S. S., Kami, K., Overduin, M., and Hilbi, H. (2009). Rab1 guanine nucleotide exchange factor SidM is a major phosphatidylinositol 4-phosphate-binding effector protein of Legionella pneumophila. J. Biol. Chem. 284, 4846–4856.

Brown, F. D., Rozelle, A. L., Yin, H. L., Balla, T., and Donaldson, J. G. (2001). Phosphatidylinositol 4,5-bisphosphate and Arf6-regulated membrane traffic. J. Cell Biol. 154, 1007–1017.

Buccione, R., Orth, J. D., and McNiven, M. A. (2004). Foot and mouth: podosomes, invadopodia and circular dorsal ruffles. Nat. Rev. Mol. Cell Biol. 5, 647–657.

Burd, C. G., and Emr, S. D. (1998). Phosphatidylinositol(3)-phosphate signaling mediated by specific binding to RING FYVE domains. Mol. Cell 2, 157–162.

Burda, P., Padilla, S. M., Sarkar, S., and Emr, S. D. (2002). Retromer function in endosome-to-Golgi retrograde transport is regulated by the yeast Vps34 PtdIns 3-kinase. J. Cell Sci. 115, 3889–3900.

Buys, S. S., and Kaplan, J. (1987). Effect of phagocytosis on receptor distribution and endocytic activity in macrophages. J. Cell. Physiol. 131, 442–449.

Campellone, K. G., and Welch, M. D. (2010). A nucleator arms race: cellular control of actin assembly. Nat. Rev. Mol. Cell Biol. 11, 237–251.

Cantalupo, G., Alifano, P., Roberti, V., Bruni, C. B., and Bucci, C. (2001). Rab-interacting lysosomal protein (RILP): the Rab7 effector required for transport to lysosomes. EMBO J. 20, 683–693.

Canton, J. (2014). Phagosome maturation in polarized macrophages. J. Leukoc. Biol. 96, 729–738.

Canton, J., Khezri, R., Glogauer, M., and Grinstein, S. (2014). Contrasting phagosome pH regulation and maturation in human M1 and M2 macrophages. Mol. Biol. Cell 25, 3330–3341.

Canton, J., Schlam, D., Breuer, C., Gütschow, M., Glogauer, M., and Grinstein, S. (2016). Calcium-sensing receptors signal constitutive macropinocytosis and facilitate the uptake of NOD2 ligands in macrophages. Nat. Commun. 7, 11284.

Champion, J. A., and Mitragotri, S. (2006). Role of target geometry in phagocytosis. Proc. Natl. Acad. Sci. 103, 4930–4934.

Charpentier, T. H., Waldo, G. L., Barrett, M. O., Huang, W., Zhang, Q., Harden, T. K., and Sondek, J. (2014). Membrane-induced allosteric control of phospholipase C-β isozymes. J. Biol. Chem. 289, 29545–29557.

Chaudhary, A., Chen, J., Gu, Q. M., Witke, W., Kwiatkowski, D. J., and Prestwich, G. D. (1998). Probing the phosphoinositide 4,5-bisphosphate binding site of human profilin I. Chem. Biol. 5, 273–281.

Chen, B.-C. et al. (2014). Lattice light-sheet microscopy: imaging molecules to embryos at high spatiotemporal resolution. Science 346, 1257998.

156

Chen, Y.-J., Hsieh, M.-Y., Chang, M. Y., Chen, H.-C., Jan, M.-S., Maa, M.-C., and Leu, T.-H. (2012). Eps8 protein facilitates phagocytosis by increasing TLR4-MyD88 protein interaction in lipopolysaccharide-stimulated macrophages. J. Biol. Chem. 287, 18806–18819.

Cheng, N., He, R., Tian, J., Dinauer, M. C., and Ye, R. D. (2007). A critical role of protein kinase C delta activation loop phosphorylation in formyl-methionyl-leucyl-phenylalanine-induced phosphorylation of p47(phox) and rapid activation of nicotinamide adenine dinucleotide phosphate oxidase. J. Immunol. Baltim. Md 1950 179, 7720–7728.

Ching, T. T., Wang, D. S., Hsu, A. L., Lu, P. J., and Chen, C. S. (1999). Identification of multiple phosphoinositide-specific phospholipases D as new regulatory enzymes for phosphatidylinositol 3,4, 5-trisphosphate. J. Biol. Chem. 274, 8611–8617.

Christoforidis, S., McBride, H. M., Burgoyne, R. D., and Zerial, M. (1999a). The Rab5 effector EEA1 is a core component of endosome docking. Nature 397, 621–625.

Christoforidis, S., Miaczynska, M., Ashman, K., Wilm, M., Zhao, L., Yip, S. C., Waterfield, M. D., Backer, J. M., and Zerial, M. (1999b). Phosphatidylinositol-3-OH kinases are Rab5 effectors. Nat. Cell Biol. 1, 249–252.

Chung, J., Nakatsu, F., Baskin, J. M., and De Camilli, P. (2015a). Plasticity of PI4KIIIα interactions at the plasma membrane. EMBO Rep. 16, 312–320.

Chung, J., Torta, F., Masai, K., Lucast, L., Czapla, H., Tanner, L. B., Narayanaswamy, P., Wenk, M. R., Nakatsu, F., and De Camilli, P. (2015b). PI4P/phosphatidylserine countertransport at ORP5- and ORP8-mediated ER-plasma membrane contacts. Science 349, 428–432.

Collins, R. F., Schreiber, A. D., Grinstein, S., and Trimble, W. S. (2002). Syntaxins 13 and 7 function at distinct steps during phagocytosis. J. Immunol. Baltim. Md 1950 169, 3250–3256.

Cooper, J. A., and Sept, D. (2008). New insights into mechanism and regulation of actin capping protein. Int. Rev. Cell Mol. Biol. 267, 183–206.

Coppolino, M. G., Dierckman, R., Loijens, J., Collins, R. F., Pouladi, M., Jongstra-Bilen, J., Schreiber, A. D., Trimble, W. S., Anderson, R., and Grinstein, S. (2002). Inhibition of phosphatidylinositol-4-phosphate 5-kinase Ialpha impairs localized actin remodeling and suppresses phagocytosis. J. Biol. Chem. 277, 43849–43857.

Cox, D., Berg, J. S., Cammer, M., Chinegwundoh, J. O., Dale, B. M., Cheney, R. E., and Greenberg, S. (2002). Myosin X is a downstream effector of PI(3)K during phagocytosis. Nat. Cell Biol. 4, 469–477.

Cox, D., Chang, P., Zhang, Q., Reddy, P. G., Bokoch, G. M., and Greenberg, S. (1997). Requirements for both Rac1 and Cdc42 in membrane ruffling and phagocytosis in leukocytes. J. Exp. Med. 186, 1487–1494.

Cox, D., Lee, D. J., Dale, B. M., Calafat, J., and Greenberg, S. (2000). A Rab11-containing rapidly recycling compartment in macrophages that promotes phagocytosis. Proc. Natl. Acad. Sci. U. S. A. 97, 680–685.

Cox, D., Tseng, C. C., Bjekic, G., and Greenberg, S. (1999a). A requirement for phosphatidylinositol 3-kinase in pseudopod extension. J. Biol. Chem. 274, 1240–1247.

157

Cox, D., Tseng, C. C., Bjekic, G., and Greenberg, S. (1999b). A requirement for phosphatidylinositol 3-kinase in pseudopod extension. J. Biol. Chem. 274, 1240–1247.

Cozier, G. E., Carlton, J., McGregor, A. H., Gleeson, P. A., Teasdale, R. D., Mellor, H., and Cullen, P. J. (2002). The phox homology (PX) domain-dependent, 3-phosphoinositide-mediated association of sorting nexin-1 with an early sorting endosomal compartment is required for its ability to regulate epidermal growth factor receptor degradation. J. Biol. Chem. 277, 48730–48736.

Craige, B., Salazar, G., and Faundez, V. (2008). Phosphatidylinositol-4-kinase type II alpha contains an AP-3-sorting motif and a kinase domain that are both required for endosome traffic. Mol. Biol. Cell 19, 1415–1426.

Crowley, M. T., Costello, P. S., Fitzer-Attas, C. J., Turner, M., Meng, F., Lowell, C., Tybulewicz, V. L., and DeFranco, A. L. (1997). A critical role for Syk in signal transduction and phagocytosis mediated by Fcgamma receptors on macrophages. J. Exp. Med. 186, 1027–1039.

Damiani, M. T., Pavarotti, M., Leiva, N., Lindsay, A. J., McCaffrey, M. W., and Colombo, M. I. (2004). Rab coupling protein associates with phagosomes and regulates recycling from the phagosomal compartment. Traffic Cph. Den. 5, 785–797.

Dart, A. E., Donnelly, S. K., Holden, D. W., Way, M., and Caron, E. (2012). Nck and Cdc42 co-operate to recruit N-WASP to promote FcγR-mediated phagocytosis. J. Cell Sci. 125, 2825–2830.

Daumke, O., Roux, A., and Haucke, V. (2014). BAR domain scaffolds in dynamin-mediated membrane fission. Cell 156, 882–892.

Dayam, R. M., Sun, C. X., Choy, C. H., Mancuso, G., Glogauer, M., and Botelho, R. J. (2017). The Lipid Kinase PIKfyve Coordinates the Neutrophil Immune Response through the Activation of the Rac GTPase. J. Immunol. Baltim. Md 1950 199, 2096–2105.

Deng, Z. et al. (2015). Tyrosine phosphatase SHP-2 mediates C-type lectin receptor-induced activation of the kinase Syk and anti-fungal TH17 responses. Nat. Immunol. 16, 642–652.

Deretic, V. et al. (2007). Phosphoinositides in phagolysosome and autophagosome biogenesis. Biochem. Soc. Symp., 141–148.

Devosse, T. et al. (2009). Formyl peptide receptor-like 2 is expressed and functional in plasmacytoid dendritic cells, tissue-specific macrophage subpopulations, and eosinophils. J. Immunol. Baltim. Md 1950 182, 4974–4984.

Di Paolo, G., and De Camilli, P. (2006). Phosphoinositides in cell regulation and membrane dynamics. Nature 443, 651–657.

Di Paolo, G., Moskowitz, H. S., Gipson, K., Wenk, M. R., Voronov, S., Obayashi, M., Flavell, R., Fitzsimonds, R. M., Ryan, T. A., and De Camilli, P. (2004). Impaired PtdIns(4,5)P2 synthesis in nerve terminals produces defects in synaptic vesicle trafficking. Nature 431, 415–422.

Dibble, C. C., and Manning, B. D. (2013). Signal integration by mTORC1 coordinates nutrient input with biosynthetic output. Nat. Cell Biol. 15, 555–564.

158

Divecha, N., Roefs, M., Halstead, J. R., D’Andrea, S., Fernandez-Borga, M., Oomen, L., Saqib, K. M., Wakelam, M. J., and D’Santos, C. (2000). Interaction of the type Ialpha PIPkinase with phospholipase D: a role for the local generation of phosphatidylinositol 4, 5-bisphosphate in the regulation of PLD2 activity. EMBO J. 19, 5440–5449.

Dodd, D. A., Worth, R. G., Rosen, M. K., Grinstein, S., van Oers, N. S. C., and Hansen, E. J. (2014). The Haemophilus ducreyi LspA1 protein inhibits phagocytosis by using a new mechanism involving activation of C-terminal Src kinase. MBio 5, e01178-01114.

Dong, R., Saheki, Y., Swarup, S., Lucast, L., Harper, J. W., and De Camilli, P. (2016). Endosome-ER Contacts Control Actin Nucleation and Retromer Function through VAP-Dependent Regulation of PI4P. Cell 166, 408–423.

Downey, G. P., Botelho, R. J., Butler, J. R., Moltyaner, Y., Chien, P., Schreiber, A. D., and Grinstein, S. (1999). Phagosomal maturation, acidification, and inhibition of bacterial growth in nonphagocytic cells transfected with FcgammaRIIA receptors. J. Biol. Chem. 274, 28436–28444.

Du, W. et al. (2016). Kinesin 1 Drives Autolysosome Tubulation. Dev. Cell 37, 326–336.

Dupré-Crochet, S., Erard, M., and Nüβe, O. (2013). ROS production in phagocytes: why, when, and where? J. Leukoc. Biol. 94, 657–670.

Elliott, M. R., and Ravichandran, K. S. (2010). Clearance of apoptotic cells: implications in health and disease. J. Cell Biol. 189, 1059–1070.

Elliott, M. R., and Ravichandran, K. S. (2016). The Dynamics of Apoptotic Cell Clearance. Dev. Cell 38, 147–160.

Epp, N., and Ungermann, C. (2013). The N-terminal domains of Vps3 and Vps8 are critical for localization and function of the CORVET tethering complex on endosomes. PloS One 8, e67307.

Fairn, G. D., and Grinstein, S. (2012). How nascent phagosomes mature to become phagolysosomes. Trends Immunol. 33, 397–405.

Fairn, G. D., Ogata, K., Botelho, R. J., Stahl, P. D., Anderson, R. A., De Camilli, P., Meyer, T., Wodak, S., and Grinstein, S. (2009). An electrostatic switch displaces phosphatidylinositol phosphate kinases from the membrane during phagocytosis. J. Cell Biol. 187, 701–714.

Falasca, M., Logan, S. K., Lehto, V. P., Baccante, G., Lemmon, M. A., and Schlessinger, J. (1998). Activation of phospholipase C gamma by PI 3-kinase-induced PH domain-mediated membrane targeting. EMBO J. 17, 414–422.

Famiglietti, S. J., Nakamura, K., and Cambier, J. C. (1999). Unique features of SHIP, SHP-1 and SHP-2 binding to FcgammaRIIb revealed by surface plasmon resonance analysis. Immunol. Lett. 68, 35–40.

Farrera, C., and Fadeel, B. (2013). Macrophage clearance of neutrophil extracellular traps is a silent process. J. Immunol. Baltim. Md 1950 191, 2647–2656.

159

Fazeli, G., Stetter, M., Lisack, J. N., and Wehman, A. M. (2018). C. elegans Blastomeres Clear the Corpse of the Second Polar Body by LC3-Associated Phagocytosis. Cell Rep. 23, 2070–2082.

Femling, J. K., Nauseef, W. M., and Weiss, J. P. (2005). Synergy between extracellular group IIA phospholipase A2 and phagocyte NADPH oxidase in digestion of phospholipids of Staphylococcus aureus ingested by human neutrophils. J. Immunol. Baltim. Md 1950 175, 4653–4661.

Ferguson, C. J., Lenk, G. M., and Meisler, M. H. (2010). PtdIns(3,5)P2 and autophagy in mouse models of neurodegeneration. Autophagy 6, 170–171.

Ferron, M., and Vacher, J. (2006). Characterization of the murine Inpp4b gene and identification of a novel isoform. Gene 376, 152–161.

von Filseck, J. M., Vanni, S., Mesmin, B., Antonny, B., and Drin, G. (2015). A phosphatidylinositol-4-phosphate powered exchange mechanism to create a lipid gradient between membranes. Nat. Commun. 6.

Fischer, K., Chatterjee, D., Torrelles, J., Brennan, P. J., Kaufmann, S. H., and Schaible, U. E. (2001). Mycobacterial lysocardiolipin is exported from phagosomes upon cleavage of cardiolipin by a macrophage-derived lysosomal phospholipase A2. J. Immunol. Baltim. Md 1950 167, 2187–2192.

Fitzer-Attas, C. J., Lowry, M., Crowley, M. T., Finn, A. J., Meng, F., DeFranco, A. L., and Lowell, C. A. (2000). Fcgamma receptor-mediated phagocytosis in macrophages lacking the Src family tyrosine kinases Hck, Fgr, and Lyn. J. Exp. Med. 191, 669–682.

Flannagan, R. S., Canton, J., Furuya, W., Glogauer, M., and Grinstein, S. (2014). The phosphatidylserine receptor TIM4 utilizes integrins as coreceptors to effect phagocytosis. Mol. Biol. Cell 25, 1511–1522.

Flannagan, R. S., and Grinstein, S. (2010). The Application of Fluorescent Probes for the Analysis of Lipid Dynamics During Phagocytosis. In: Live Cell Imaging: Methods and Protocols, ed. D. B. Papkovsky, Totowa, NJ: Humana Press, 121–134.

Flannagan, R. S., Harrison, R. E., Yip, C. M., Jaqaman, K., and Grinstein, S. (2010). Dynamic macrophage “probing” is required for the efficient capture of phagocytic targets. J. Cell Biol. 191, 1205–1218.

Flannagan, R. S., Jaumouillé, V., and Grinstein, S. (2012). The cell biology of phagocytosis. Annu. Rev. Pathol. 7, 61–98.

Foti, M., Audhya, A., and Emr, S. D. (2001). Sac1 lipid phosphatase and Stt4 phosphatidylinositol 4-kinase regulate a pool of phosphatidylinositol 4-phosphate that functions in the control of the actin cytoskeleton and vacuole morphology. Mol. Biol. Cell 12, 2396–2411.

Fratti, R. A., Backer, J. M., Gruenberg, J., Corvera, S., and Deretic, V. (2001a). Role of phosphatidylinositol 3-kinase and Rab5 effectors in phagosomal biogenesis and mycobacterial phagosome maturation arrest. J. Cell Biol. 154, 631–644.

160

Fratti, R. A., Backer, J. M., Gruenberg, J., Corvera, S., and Deretic, V. (2001b). Role of phosphatidylinositol 3-kinase and Rab5 effectors in phagosomal biogenesis and mycobacterial phagosome maturation arrest. J. Cell Biol. 154, 631–644.

Fratti, R. A., Chua, J., Vergne, I., and Deretic, V. (2003). Mycobacterium tuberculosis glycosylated phosphatidylinositol causes phagosome maturation arrest. Proc. Natl. Acad. Sci. U. S. A. 100, 5437–5442.

Freedman, T. S., Tan, Y. X., Skrzypczynska, K. M., Manz, B. N., Sjaastad, F. V., Goodridge, H. S., Lowell, C. A., and Weiss, A. (2015). LynA regulates an inflammation-sensitive signaling checkpoint in macrophages. ELife 4.

Freeman, S. A. et al. (2015). Toll-like receptor ligands sensitize B-cell receptor signalling by reducing actin-dependent spatial confinement of the receptor. Nat. Commun. 6, 6168.

Freeman, S. A., Goyette, J., Furuya, W., Woods, E. C., Bertozzi, C. R., Bergmeier, W., Hinz, B., van der Merwe, P. A., Das, R., and Grinstein, S. (2016). Integrins Form an Expanding Diffusional Barrier that Coordinates Phagocytosis. Cell 164, 128–140.

Freitas, R. A. (2000). Andrew Nanomedicine, Volume 1: Basic Capabilities, by Robert A. Freitas., Landes Bioscience.

Friedland, N., Liou, H.-L., Lobel, P., and Stock, A. M. (2003). Structure of a cholesterol-binding protein deficient in Niemann-Pick type C2 disease. Proc. Natl. Acad. Sci. U. S. A. 100, 2512–2517.

Frischknecht, F., and Way, M. (2001). Surfing pathogens and the lessons learned for actin polymerization. Trends Cell Biol. 11, 30–38.

Fu, Y., and Galán, J. E. (1999). A salmonella protein antagonizes Rac-1 and Cdc42 to mediate host-cell recovery after bacterial invasion. Nature 401, 293–297.

Fujiwara, T. K. et al. (2016). Confined diffusion of transmembrane proteins and lipids induced by the same actin meshwork lining the plasma membrane. Mol. Biol. Cell 27, 1101–1119.

Ganesan, L. P., Fang, H., Marsh, C. B., and Tridandapani, S. (2003). The protein-tyrosine phosphatase SHP-1 associates with the phosphorylated immunoreceptor tyrosine-based activation motif of Fc gamma RIIa to modulate signaling events in myeloid cells. J. Biol. Chem. 278, 35710–35717.

Garin, J., Diez, R., Kieffer, S., Dermine, J. F., Duclos, S., Gagnon, E., Sadoul, R., Rondeau, C., and Desjardins, M. (2001). The phagosome proteome: insight into phagosome functions. J. Cell Biol. 152, 165–180.

Getahun, A., and Cambier, J. C. (2015). Of ITIMs, ITAMs, and ITAMis: revisiting immunoglobulin Fc receptor signaling. Immunol. Rev. 268, 66–73.

Gillooly, D. J., Morrow, I. C., Lindsay, M., Gould, R., Bryant, N. J., Gaullier, J. M., Parton, R. G., and Stenmark, H. (2000). Localization of phosphatidylinositol 3-phosphate in yeast and mammalian cells. EMBO J. 19, 4577–4588.

Gillooly, D. J., Simonsen, A., and Stenmark, H. (2001). Phosphoinositides and phagocytosis. J. Cell Biol. 155, 15–18.

161

Godi, A., Pertile, P., Meyers, R., Marra, P., Di Tullio, G., Iurisci, C., Luini, A., Corda, D., and De Matteis, M. A. (1999). ARF mediates recruitment of PtdIns-4-OH kinase-beta and stimulates synthesis of PtdIns(4,5)P2 on the Golgi complex. Nat. Cell Biol. 1, 280–287.

Gold, E. S., Morrissette, N. S., Underhill, D. M., Guo, J., Bassetti, M., and Aderem, A. (2000). Amphiphysin IIm, a novel amphiphysin II isoform, is required for macrophage phagocytosis. Immunity 12, 285–292.

Gold, E. S., Underhill, D. M., Morrissette, N. S., Guo, J., McNiven, M. A., and Aderem, A. (1999). Dynamin 2 is required for phagocytosis in macrophages. J. Exp. Med. 190, 1849–1856.

Goley, E. D., Rodenbusch, S. E., Martin, A. C., and Welch, M. D. (2004). Critical conformational changes in the Arp2/3 complex are induced by nucleotide and nucleation promoting factor. Mol. Cell 16, 269–279.

Goodridge, H. S. et al. (2011). Activation of the innate immune receptor Dectin-1 upon formation of a “phagocytic synapse.” Nature 472, 471–475.

Gorbatyuk, V. Y., Nosworthy, N. J., Robson, S. A., Bains, N. P. S., Maciejewski, M. W., Dos Remedios, C. G., and King, G. F. (2006). Mapping the phosphoinositide-binding site on chick cofilin explains how PIP2 regulates the cofilin-actin interaction. Mol. Cell 24, 511–522.

Gordon, A. H., Hart, P. D., and Young, M. R. (1980). Ammonia inhibits phagosome-lysosome fusion in macrophages. Nature 286, 79–80.

Gottlieb, Y., Topaz, O., Cohen, L. A., Yakov, L. D., Haber, T., Morgenstern, A., Weiss, A., Chait Berman, K., Fibach, E., and Meyron-Holtz, E. G. (2012). Physiologically aged red blood cells undergo erythrophagocytosis in vivo but not in vitro. Haematologica 97, 994–1002.

Graham, T. R., and Burd, C. G. (2011). Coordination of Golgi functions by phosphatidylinositol 4-kinases. Trends Cell Biol. 21, 113–121.

Gray, M., and Botelho, R. J. (2017). Phagocytosis: Hungry, Hungry Cells. Methods Mol. Biol. Clifton NJ 1519, 1–16.

Gude, D. R., Alvarez, S. E., Paugh, S. W., Mitra, P., Yu, J., Griffiths, R., Barbour, S. E., Milstien, S., and Spiegel, S. (2008). Apoptosis induces expression of sphingosine kinase 1 to release sphingosine-1-phosphate as a “come-and-get-me” signal. FASEB J. Off. Publ. Fed. Am. Soc. Exp. Biol. 22, 2629–2638.

Guilliams, M., Bruhns, P., Saeys, Y., Hammad, H., and Lambrecht, B. N. (2014). The function of Fcγ receptors in dendritic cells and macrophages. Nat. Rev. Immunol. 14, 94–108.

Guo, F., Ding, Y., Caberoy, N., Alvarado, G., Wang, F., Chen, R., and Li, W. (2015). ABCF1 extrinsically regulates retinal pigment epithelial cell phagocytosis. Mol. Biol. Cell 26, 2311–2320.

Guo, S., Stolz, L. E., Lemrow, S. M., and York, J. D. (1999). SAC1-like domains of yeast SAC1, INP52, and INP53 and of human synaptojanin encode polyphosphoinositide phosphatases. J. Biol. Chem. 274, 12990–12995.

Hackam, D. J., Rotstein, O. D., Zhang, W. J., Demaurex, N., Woodside, M., Tsai, O., and Grinstein, S. (1997). Regulation of phagosomal acidification. Differential targeting of Na+/H+

162

exchangers, Na+/K+-ATPases, and vacuolar-type H+-atpases. J. Biol. Chem. 272, 29810–29820.

Hall, A. B., Gakidis, M. A. M., Glogauer, M., Wilsbacher, J. L., Gao, S., Swat, W., and Brugge, J. S. (2006). Requirements for Vav guanine nucleotide exchange factors and Rho GTPases in FcgammaR- and complement-mediated phagocytosis. Immunity 24, 305–316.

Ham, H., Sreelatha, A., and Orth, K. (2011). Manipulation of host membranes by bacterial effectors. Nat. Rev. Microbiol. 9, 635–646.

Hamada, K., Shimizu, T., Matsui, T., Tsukita, S., and Hakoshima, T. (2000). Structural basis of the membrane-targeting and unmasking mechanisms of the radixin FERM domain. EMBO J. 19, 4449–4462.

Hammond, G. R. V., and Balla, T. (2015). Polyphosphoinositide binding domains: Key to inositol lipid biology. Biochim. Biophys. Acta 1851, 746–758.

Hammond, G. R. V., Fischer, M. J., Anderson, K. E., Holdich, J., Koteci, A., Balla, T., and Irvine, R. F. (2012). PI4P and PI(4,5)P2 are essential but independent lipid determinants of membrane identity. Science 337, 727–730.

Hammond, G. R. V., Machner, M. P., and Balla, T. (2014). A novel probe for phosphatidylinositol 4-phosphate reveals multiple pools beyond the Golgi. J. Cell Biol. 205, 113–126.

Hammond, G. R. V., Schiavo, G., and Irvine, R. F. (2009). Immunocytochemical techniques reveal multiple, distinct cellular pools of PtdIns4P and PtdIns(4,5)P(2). Biochem. J. 422, 23–35.

Harrison, R. E., Bucci, C., Vieira, O. V., Schroer, T. A., and Grinstein, S. (2003). Phagosomes fuse with late endosomes and/or lysosomes by extension of membrane protrusions along microtubules: role of Rab7 and RILP. Mol. Cell. Biol. 23, 6494–6506.

He, R., Nanamori, M., Sang, H., Yin, H., Dinauer, M. C., and Ye, R. D. (2004). Reconstitution of chemotactic peptide-induced nicotinamide adenine dinucleotide phosphate (reduced) oxidase activation in transgenic COS-phox cells. J. Immunol. Baltim. Md 1950 173, 7462–7470.

Heit, B., Liu, L., Colarusso, P., Puri, K. D., and Kubes, P. (2008). PI3K accelerates, but is not required for, neutrophil chemotaxis to fMLP. J. Cell Sci. 121, 205–214.

Herre, J., Marshall, A. S. J., Caron, E., Edwards, A. D., Williams, D. L., Schweighoffer, E., Tybulewicz, V., Reis e Sousa, C., Gordon, S., and Brown, G. D. (2004). Dectin-1 uses novel mechanisms for yeast phagocytosis in macrophages. Blood 104, 4038–4045.

Hierro, A., Rojas, A. L., Rojas, R., Murthy, N., Effantin, G., Kajava, A. V., Steven, A. C., Bonifacino, J. S., and Hurley, J. H. (2007). Functional architecture of the retromer cargo-recognition complex. Nature 449, 1063–1067.

Hiraoka, M., Abe, A., Lu, Y., Yang, K., Han, X., Gross, R. W., and Shayman, J. A. (2006). Lysosomal phospholipase A2 and phospholipidosis. Mol. Cell. Biol. 26, 6139–6148.

Honda, A. et al. (1999). Phosphatidylinositol 4-phosphate 5-kinase alpha is a downstream effector of the small G protein ARF6 in membrane ruffle formation. Cell 99, 521–532.

163

Horiuchi, H. et al. (1997). A novel Rab5 GDP/GTP exchange factor complexed to Rabaptin-5 links nucleotide exchange to effector recruitment and function. Cell 90, 1149–1159.

Hsu, C.-L. et al. (2012). Equilibrative nucleoside transporter 3 deficiency perturbs lysosome function and macrophage homeostasis. Science 335, 89–92.

Hsu, F., Hu, F., and Mao, Y. (2015). Spatiotemporal control of phosphatidylinositol 4-phosphate by Sac2 regulates endocytic recycling. J. Cell Biol. 209, 97–110.

Huang, S. C.-C. et al. (2014). Cell-intrinsic lysosomal lipolysis is essential for alternative activation of macrophages. Nat. Immunol. 15, 846–855.

Humphries, A. C., Donnelly, S. K., and Way, M. (2014). Cdc42 and the Rho GEF intersectin-1 collaborate with Nck to promote N-WASP-dependent actin polymerisation. J. Cell Sci. 127, 673–685.

Huynh, K. K., Eskelinen, E.-L., Scott, C. C., Malevanets, A., Saftig, P., and Grinstein, S. (2007). LAMP proteins are required for fusion of lysosomes with phagosomes. EMBO J. 26, 313–324.

Huynh, K. K., Gershenzon, E., and Grinstein, S. (2008). Cholesterol accumulation by macrophages impairs phagosome maturation. J. Biol. Chem. 283, 35745–35755.

Ikonomov, O. C., Sbrissa, D., Mlak, K., Kanzaki, M., Pessin, J., and Shisheva, A. (2002). Functional dissection of lipid and protein kinase signals of PIKfyve reveals the role of PtdIns 3,5-P2 production for endomembrane integrity. J. Biol. Chem. 277, 9206–9211.

Indik, Z., Kelly, C., Chien, P., Levinson, A. I., and Schreiber, A. D. (1991). Human Fc gamma RII, in the absence of other Fc gamma receptors, mediates a phagocytic signal. J. Clin. Invest. 88, 1766–1771.

Infante, R. E., Wang, M. L., Radhakrishnan, A., Kwon, H. J., Brown, M. S., and Goldstein, J. L. (2008). NPC2 facilitates bidirectional transfer of cholesterol between NPC1 and lipid bilayers, a step in cholesterol egress from lysosomes. Proc. Natl. Acad. Sci. U. S. A. 105, 15287–15292.

Innocenti, M. et al. (2005). Abi1 regulates the activity of N-WASP and WAVE in distinct actin-based processes. Nat. Cell Biol. 7, 969–976.

Isogai, T., van der Kammen, R., Leyton-Puig, D., Kedziora, K. M., Jalink, K., and Innocenti, M. (2015). Initiation of lamellipodia and ruffles involves cooperation between mDia1 and the Arp2/3 complex. J. Cell Sci. 128, 3796–3810.

Itoh, T., Erdmann, K. S., Roux, A., Habermann, B., Werner, H., and De Camilli, P. (2005). Dynamin and the actin cytoskeleton cooperatively regulate plasma membrane invagination by BAR and F-BAR proteins. Dev. Cell 9, 791–804.

Iyer, S. S., Barton, J. A., Bourgoin, S., and Kusner, D. J. (2004). Phospholipases D1 and D2 coordinately regulate macrophage phagocytosis. J. Immunol. Baltim. Md 1950 173, 2615–2623.

Jaconi, M. E., Lew, D. P., Carpentier, J. L., Magnusson, K. E., Sjögren, M., and Stendahl, O. (1990). Cytosolic free calcium elevation mediates the phagosome-lysosome fusion during phagocytosis in human neutrophils. J. Cell Biol. 110, 1555–1564.

164

Jankowski, A., Scott, C. C., and Grinstein, S. (2002). Determinants of the phagosomal pH in neutrophils. J. Biol. Chem. 277, 6059–6066.

Janmey, P. A., and Stossel, T. P. (1987). Modulation of gelsolin function by phosphatidylinositol 4,5-bisphosphate. Nature 325, 362–364.

Jaqaman, K., Kuwata, H., Touret, N., Collins, R., Trimble, W. S., Danuser, G., and Grinstein, S. (2011). Cytoskeletal control of CD36 diffusion promotes its receptor and signaling function. Cell 146, 593–606.

Jaumouillé, V., Farkash, Y., Jaqaman, K., Das, R., Lowell, C. A., and Grinstein, S. (2014). Actin cytoskeleton reorganization by Syk regulates Fcγ receptor responsiveness by increasing its lateral mobility and clustering. Dev. Cell 29, 534–546.

Jaumouillé, V., and Grinstein, S. (2011). Receptor mobility, the cytoskeleton, and particle binding during phagocytosis. Curr. Opin. Cell Biol. 23, 22–29.

Jefferies, H. B. J. et al. (2008). A selective PIKfyve inhibitor blocks PtdIns(3,5)P(2) production and disrupts endomembrane transport and retroviral budding. EMBO Rep. 9, 164–170.

Jeschke, A. et al. (2015). Phosphatidylinositol 4-phosphate and phosphatidylinositol 3-phosphate regulate phagolysosome biogenesis. Proc. Natl. Acad. Sci. U. S. A. 112, 4636–4641.

Johansson, M., Bocher, V., Lehto, M., Chinetti, G., Kuismanen, E., Ehnholm, C., Staels, B., and Olkkonen, V. M. (2003). The two variants of oxysterol binding protein-related protein-1 display different tissue expression patterns, have different intracellular localization, and are functionally distinct. Mol. Biol. Cell 14, 903–915.

Johansson, M., Lehto, M., Tanhuanpää, K., Cover, T. L., and Olkkonen, V. M. (2005). The oxysterol-binding protein homologue ORP1L interacts with Rab7 and alters functional properties of late endocytic compartments. Mol. Biol. Cell 16, 5480–5492.

Johansson, M., Rocha, N., Zwart, W., Jordens, I., Janssen, L., Kuijl, C., Olkkonen, V. M., and Neefjes, J. (2007). Activation of endosomal dynein motors by stepwise assembly of Rab7-RILP-p150Glued, ORP1L, and the receptor betalll spectrin. J. Cell Biol. 176, 459–471.

Johnson, D. E., Ostrowski, P., Jaumouillé, V., and Grinstein, S. (2016). The position of lysosomes within the cell determines their luminal pH. J. Cell Biol. 212, 677–692.

Johnson, S. A., Pleiman, C. M., Pao, L., Schneringer, J., Hippen, K., and Cambier, J. C. (1995). Phosphorylated immunoreceptor signaling motifs (ITAMs) exhibit unique abilities to bind and activate Lyn and Syk tyrosine kinases. J. Immunol. Baltim. Md 1950 155, 4596–4603.

Jordens, I., Fernandez-Borja, M., Marsman, M., Dusseljee, S., Janssen, L., Calafat, J., Janssen, H., Wubbolts, R., and Neefjes, J. (2001). The Rab7 effector protein RILP controls lysosomal transport by inducing the recruitment of dynein-dynactin motors. Curr. Biol. CB 11, 1680–1685.

Jović, M., Kean, M. J., Szentpetery, Z., Polevoy, G., Gingras, A.-C., Brill, J. A., and Balla, T. (2012). Two phosphatidylinositol 4-kinases control lysosomal delivery of the Gaucher disease enzyme, β-glucocerebrosidase. Mol. Biol. Cell 23, 1533–1545.

Kaksonen, M., Toret, C. P., and Drubin, D. G. (2006). Harnessing actin dynamics for clathrin-mediated endocytosis. Nat. Rev. Mol. Cell Biol. 7, 404–414.

165

Kamen, L. A., Levinsohn, J., and Swanson, J. A. (2007). Differential association of phosphatidylinositol 3-kinase, SHIP-1, and PTEN with forming phagosomes. Mol. Biol. Cell 18, 2463–2472.

Kanchanawong, P., Shtengel, G., Pasapera, A. M., Ramko, E. B., Davidson, M. W., Hess, H. F., and Waterman, C. M. (2010). Nanoscale architecture of integrin-based cell adhesions. Nature 468, 580–584.

van der Kant, R., Fish, A., Janssen, L., Janssen, H., Krom, S., Ho, N., Brummelkamp, T., Carette, J., Rocha, N., and Neefjes, J. (2013a). Late endosomal transport and tethering are coupled processes controlled by RILP and the cholesterol sensor ORP1L. J. Cell Sci. 126, 3462–3474.

van der Kant, R., Zondervan, I., Janssen, L., and Neefjes, J. (2013b). Cholesterol-binding molecules MLN64 and ORP1L mark distinct late endosomes with transporters ABCA3 and NPC1. J. Lipid Res. 54, 2153–2165.

Kawane, K., Fukuyama, H., Kondoh, G., Takeda, J., Ohsawa, Y., Uchiyama, Y., and Nagata, S. (2001). Requirement of DNase II for definitive erythropoiesis in the mouse fetal liver. Science 292, 1546–1549.

Ketel, K., Krauss, M., Nicot, A.-S., Puchkov, D., Wieffer, M., Müller, R., Subramanian, D., Schultz, C., Laporte, J., and Haucke, V. (2016). A phosphoinositide conversion mechanism for exit from endosomes. Nature 529, 408–412.

Kheir, W. A., Gevrey, J.-C., Yamaguchi, H., Isaac, B., and Cox, D. (2005). A WAVE2-Abi1 complex mediates CSF-1-induced F-actin-rich membrane protrusions and migration in macrophages. J. Cell Sci. 118, 5369–5379.

Kiefer, F., Brumell, J., Al-Alawi, N., Latour, S., Cheng, A., Veillette, A., Grinstein, S., and Pawson, T. (1998). The Syk protein tyrosine kinase is essential for Fcgamma receptor signaling in macrophages and neutrophils. Mol. Cell. Biol. 18, 4209–4220.

Kinchen, J. M., Doukoumetzidis, K., Almendinger, J., Stergiou, L., Tosello-Trampont, A., Sifri, C. D., Hengartner, M. O., and Ravichandran, K. S. (2008). A pathway for phagosome maturation during engulfment of apoptotic cells. Nat. Cell Biol. 10, 556–566.

Kinchen, J. M., and Ravichandran, K. S. (2008). Phagosome maturation: going through the acid test. Nat. Rev. Mol. Cell Biol. 9, 781–795.

Kinchen, J. M., and Ravichandran, K. S. (2010). Identification of two evolutionarily conserved genes regulating processing of engulfed apoptotic cells. Nature 464, 778–782.

Kitano, M., Nakaya, M., Nakamura, T., Nagata, S., and Matsuda, M. (2008). Imaging of Rab5 activity identifies essential regulators for phagosome maturation. Nature 453, 241–245.

Koh, A. L. Y., Sun, C. X., Zhu, F., and Glogauer, M. (2005). The role of Rac1 and Rac2 in bacterial killing. Cell. Immunol. 235, 92–97.

Kohyama, M., Ise, W., Edelson, B. T., Wilker, P. R., Hildner, K., Mejia, C., Frazier, W. A., Murphy, T. L., and Murphy, K. M. (2009). Role for Spi-C in the development of red pulp macrophages and splenic iron homeostasis. Nature 457, 318–321.

166

Koivusalo, M., Welch, C., Hayashi, H., Scott, C. C., Kim, M., Alexander, T., Touret, N., Hahn, K. M., and Grinstein, S. (2010). Amiloride inhibits macropinocytosis by lowering submembranous pH and preventing Rac1 and Cdc42 signaling. J. Cell Biol. 188, 547–563.

Kolter, T., and Sandhoff, K. (2010). Lysosomal degradation of membrane lipids. FEBS Lett. 584, 1700–1712.

Kong, L., and Ge, B.-X. (2008). MyD88-independent activation of a novel actin-Cdc42/Rac pathway is required for Toll-like receptor-stimulated phagocytosis. Cell Res. 18, 745–755.

Krajcovic, M., Krishna, S., Akkari, L., Joyce, J. A., and Overholtzer, M. (2013). mTOR regulates phagosome and entotic vacuole fission. Mol. Biol. Cell 24, 3736–3745.

Kreishman-Deitrick, M., Goley, E. D., Burdine, L., Denison, C., Egile, C., Li, R., Murali, N., Kodadek, T. J., Welch, M. D., and Rosen, M. K. (2005). NMR analyses of the activation of the Arp2/3 complex by neuronal Wiskott-Aldrich syndrome protein. Biochemistry (Mosc.) 44, 15247–15256.

Krieser, R. J., MacLea, K. S., Longnecker, D. S., Fields, J. L., Fiering, S., and Eastman, A. (2002). Deoxyribonuclease IIalpha is required during the phagocytic phase of apoptosis and its loss causes perinatal lethality. Cell Death Differ. 9, 956–962.

Kusner, D. J., Hall, C. F., and Jackson, S. (1999). Fc gamma receptor-mediated activation of phospholipase D regulates macrophage phagocytosis of IgG-opsonized particles. J. Immunol. Baltim. Md 1950 162, 2266–2274.

Kusumi, A., Nakada, C., Ritchie, K., Murase, K., Suzuki, K., Murakoshi, H., Kasai, R. S., Kondo, J., and Fujiwara, T. (2005). Paradigm shift of the plasma membrane concept from the two-dimensional continuum fluid to the partitioned fluid: high-speed single-molecule tracking of membrane molecules. Annu. Rev. Biophys. Biomol. Struct. 34, 351–378.

Kusumi, A., Tsunoyama, T. A., Hirosawa, K. M., Kasai, R. S., and Fujiwara, T. K. (2014). Tracking single molecules at work in living cells. Nat. Chem. Biol. 10, 524–532.

Kutateladze, T. G. (2010). Translation of the phosphoinositide code by PI effectors. Nat. Chem. Biol. 6, 507–513.

Kwiatkowska, K., and Sobota, A. (1999). Signaling pathways in phagocytosis. BioEssays News Rev. Mol. Cell. Dev. Biol. 21, 422–431.

de Lartigue, J., Polson, H., Feldman, M., Shokat, K., Tooze, S. A., Urbé, S., and Clague, M. J. (2009). PIKfyve regulation of endosome-linked pathways. Traffic Cph. Den. 10, 883–893.

Lawe, D. C., Chawla, A., Merithew, E., Dumas, J., Carrington, W., Fogarty, K., Lifshitz, L., Tuft, R., Lambright, D., and Corvera, S. (2002). Sequential roles for phosphatidylinositol 3-phosphate and Rab5 in tethering and fusion of early endosomes via their interaction with EEA1. J. Biol. Chem. 277, 8611–8617.

Lee, C.-Y., Herant, M., and Heinrich, V. (2011). Target-specific mechanics of phagocytosis: protrusive neutrophil response to zymosan differs from the uptake of antibody-tagged pathogens. J. Cell Sci. 124, 1106–1114.

167

Lee, W. L., Kim, M.-K., Schreiber, A. D., and Grinstein, S. (2005). Role of ubiquitin and proteasomes in phagosome maturation. Mol. Biol. Cell 16, 2077–2090.

Lee, Y., Song, S.-M., Park, H. S., Kim, S., Koh, E.-H., Choi, M. S., and Choi, M.-U. (2002). Elevation of oleate-activated phospholipase D activity during thymic atrophy. Immunology 107, 435–443.

Lees, J. A. et al. (2017). Architecture of the human PI4KIIIα lipid kinase complex. Proc. Natl. Acad. Sci. U. S. A. 114, 13720–13725.

Lemmon, M. A. (2003). Phosphoinositide recognition domains. Traffic Cph. Den. 4, 201–213.

Levin, R., Grinstein, S., and Canton, J. (2016). The life cycle of phagosomes: formation, maturation, and resolution. Immunol. Rev. 273, 156–179.

Levin, R., Grinstein, S., and Schlam, D. (2015). Phosphoinositides in phagocytosis and macropinocytosis. Biochim. Biophys. Acta 1851, 805–823.

Levin, R., Hammond, G. R. V., Balla, T., De Camilli, P., Fairn, G. D., and Grinstein, S. (2017). Multiphasic dynamics of phosphatidylinositol 4-phosphate during phagocytosis. Mol. Biol. Cell 28, 128–140.

Lew, E. D., Oh, J., Burrola, P. G., Lax, I., Zag??rska, A., Trav??s, P. G., Schlessinger, J., and Lemke, G. (2014). Differential TAM receptor-ligand-phospholipid interactions delimit differential TAM bioactivities. ELife 3, 1–23.

Liao, F., Shin, H. S., and Rhee, S. G. (1992). Tyrosine phosphorylation of phospholipase C-gamma 1 induced by cross-linking of the high-affinity or low-affinity Fc receptor for IgG in U937 cells. Proc. Natl. Acad. Sci. U. S. A. 89, 3659–3663.

Linke, T., Wilkening, G., Lansmann, S., Moczall, H., Bartelsen, O., Weisgerber, J., and Sandhoff, K. (2001). Stimulation of acid sphingomyelinase activity by lysosomal lipids and sphingolipid activator proteins. Biol. Chem. 382, 283–290.

Linkner, J., Witte, G., Zhao, H., Junemann, A., Nordholz, B., Runge-Wollmann, P., Lappalainen, P., and Faix, J. (2014). The inverse BAR domain protein IBARa drives membrane remodeling to control osmoregulation, phagocytosis and cytokinesis. J. Cell Sci. 127, 1279–1292.

Lippé, R., Miaczynska, M., Rybin, V., Runge, A., and Zerial, M. (2001). Functional synergy between Rab5 effector Rabaptin-5 and exchange factor Rabex-5 when physically associated in a complex. Mol. Biol. Cell 12, 2219–2228.

Liscum, L. (2000). Niemann-Pick type C mutations cause lipid traffic jam. Traffic Cph. Den. 1, 218–225.

Lloyd-Evans, E., Morgan, A. J., He, X., Smith, D. A., Elliot-Smith, E., Sillence, D. J., Churchill, G. C., Schuchman, E. H., Galione, A., and Platt, F. M. (2008). Niemann-Pick disease type C1 is a sphingosine storage disease that causes deregulation of lysosomal calcium. Nat. Med. 14, 1247–1255.

Lobingier, B. T., and Merz, A. J. (2012). Sec1/Munc18 protein Vps33 binds to SNARE domains and the quaternary SNARE complex. Mol. Biol. Cell 23, 4611–4622.

168

Loewen, C. J. R., and Levine, T. P. (2005). A highly conserved binding site in vesicle-associated membrane protein-associated protein (VAP) for the FFAT motif of lipid-binding proteins. J. Biol. Chem. 280, 14097–14104.

Loewen, C. J. R., Roy, A., and Levine, T. P. (2003). A conserved ER targeting motif in three families of lipid binding proteins and in Opi1p binds VAP. EMBO J. 22, 2025–2035.

Loovers, H. M., Kortholt, A., de Groote, H., Whitty, L., Nussbaum, R. L., and van Haastert, P. J. M. (2007). Regulation of phagocytosis in Dictyostelium by the inositol 5-phosphatase OCRL homolog Dd5P4. Traffic Cph. Den. 8, 618–628.

Lorenzi, R., Brickell, P. M., Katz, D. R., Kinnon, C., and Thrasher, A. J. (2000). Wiskott-Aldrich syndrome protein is necessary for efficient IgG-mediated phagocytosis. Blood 95, 2943–2946.

Lorenzo, O., Urbé, S., and Clague, M. J. (2006). Systematic analysis of myotubularins: heteromeric interactions, subcellular localisation and endosome related functions. J. Cell Sci. 119, 2953–2959.

Lu, S. M., Grinstein, S., and Fairn, G. D. (2017). Phagocytosis and Phagosomes. 1519, 79–91.

Lucin, K. M. et al. (2013). Microglial beclin 1 regulates retromer trafficking and phagocytosis and is impaired in Alzheimer’s disease. Neuron 79, 873–886.

Lukacs, G. L., Rotstein, O. D., and Grinstein, S. (1991). Determinants of the phagosomal pH in macrophages. In situ assessment of vacuolar H(+)-ATPase activity, counterion conductance, and H+ “leak.” J. Biol. Chem. 266, 24540–24548.

Lundquist, M. R. et al. (2018). Phosphatidylinositol-5-Phosphate 4-Kinases Regulate Cellular Lipid Metabolism By Facilitating Autophagy. Mol. Cell 70, 531–544.e9.

Luzio, J. P., Pryor, P. R., and Bright, N. A. (2007). Lysosomes: fusion and function. Nat. Rev. Mol. Cell Biol. 8, 622–632.

MacDougall, L. K., Domin, J., and Waterfield, M. D. (1995). A family of phosphoinositide 3-kinases in Drosophila identifies a new mediator of signal transduction. Curr. Biol. CB 5, 1404–1415.

Maeda, K., Anand, K., Chiapparino, A., Kumar, A., Poletto, M., Kaksonen, M., and Gavin, A.-C. (2013). Interactome map uncovers phosphatidylserine transport by oxysterol-binding proteins. Nature 501, 257–261.

Maehama, T., and Dixon, J. E. (1998). The tumor suppressor, PTEN/MMAC1, dephosphorylates the lipid second messenger, phosphatidylinositol 3,4,5-trisphosphate. J. Biol. Chem. 273, 13375–13378.

Malbec, O., Fong, D. C., Turner, M., Tybulewicz, V. L., Cambier, J. C., Fridman, W. H., and Daëron, M. (1998). Fc epsilon receptor I-associated lyn-dependent phosphorylation of Fc gamma receptor IIB during negative regulation of mast cell activation. J. Immunol. Baltim. Md 1950 160, 1647–1658.

Mantegazza, A. R., Guttentag, S. H., El-Benna, J., Sasai, M., Iwasaki, A., Shen, H., Laufer, T. M., and Marks, M. S. (2012). Adaptor protein-3 in dendritic cells facilitates phagosomal toll-like receptor signaling and antigen presentation to CD4(+) T cells. Immunity 36, 782–794.

169

Mantegazza, A. R., Magalhaes, J. G., Amigorena, S., and Marks, M. S. (2013). Presentation of phagocytosed antigens by MHC class I and II. Traffic Cph. Den. 14, 135–152.

Mantegazza, A. R., Savina, A., Vermeulen, M., Pérez, L., Geffner, J., Hermine, O., Rosenzweig, S. D., Faure, F., and Amigorena, S. (2008). NADPH oxidase controls phagosomal pH and antigen cross-presentation in human dendritic cells. Blood 112, 4712–4722.

Mantegazza, A. R., Zajac, A. L., Twelvetrees, A., Holzbaur, E. L. F., Amigorena, S., and Marks, M. S. (2014). TLR-dependent phagosome tubulation in dendritic cells promotes phagosome cross-talk to optimize MHC-II antigen presentation. Proc. Natl. Acad. Sci. U. S. A. 111, 15508–15513.

Manwani, D., and Bieker, J. J. (2008). The erythroblastic island. Curr. Top. Dev. Biol. 82, 23–53.

Mao, Y. S. et al. (2009). Essential and unique roles of PIP5K-gamma and -alpha in Fcgamma receptor-mediated phagocytosis. J. Cell Biol. 184, 281–296.

Marie-Anaïs, F., Mazzolini, J., Herit, F., and Niedergang, F. (2016). Dynamin-Actin Cross Talk Contributes to Phagosome Formation and Closure. Traffic Cph. Den. 17, 487–499.

Marion, S. et al. (2012). The NF-κB signaling protein Bcl10 regulates actin dynamics by controlling AP1 and OCRL-bearing vesicles. Dev. Cell 23, 954–967.

Marquer, C., Tian, H., Yi, J., Bastien, J., Dall’Armi, C., Yang-Klingler, Y., Zhou, B., Chan, R. B., and Di Paolo, G. (2016). Arf6 controls retromer traffic and intracellular cholesterol distribution via a phosphoinositide-based mechanism. Nat. Commun. 7, 11919.

Marshall, J. G., Booth, J. W., Stambolic, V., Mak, T., Balla, T., Schreiber, A. D., Meyer, T., and Grinstein, S. (2001). Restricted accumulation of phosphatidylinositol 3-kinase products in a plasmalemmal subdomain during Fc gamma receptor-mediated phagocytosis. J. Cell Biol. 153, 1369–1380.

Martel, V., Racaud-Sultan, C., Dupe, S., Marie, C., Paulhe, F., Galmiche, A., Block, M. R., and Albiges-Rizo, C. (2001). Conformation, localization, and integrin binding of talin depend on its interaction with phosphoinositides. J. Biol. Chem. 276, 21217–21227.

Martin, G. S. (2001). The hunting of the Src. Nat. Rev. Mol. Cell Biol. 2, 467–475.

Matsuo, H. et al. (2004). Role of LBPA and Alix in multivesicular liposome formation and endosome organization. Science 303, 531–534.

Mattila, P. K., Feest, C., Depoil, D., Treanor, B., Montaner, B., Otipoby, K. L., Carter, R., Justement, L. B., Bruckbauer, A., and Batista, F. D. (2013). The actin and tetraspanin networks organize receptor nanoclusters to regulate B cell receptor-mediated signaling. Immunity 38, 461–474.

May, R. C., Caron, E., Hall, A., and Machesky, L. M. (2000). Involvement of the Arp2/3 complex in phagocytosis mediated by FcγR or CR3. Nat. Cell Biol. 2, 246–248.

McBride, H. M., Rybin, V., Murphy, C., Giner, A., Teasdale, R., and Zerial, M. (1999). Oligomeric complexes link Rab5 effectors with NSF and drive membrane fusion via interactions between EEA1 and syntaxin 13. Cell 98, 377–386.

170

McCartney, A. J., Zhang, Y., and Weisman, L. S. (2014). Phosphatidylinositol 3,5-bisphosphate: low abundance, high significance. BioEssays News Rev. Mol. Cell. Dev. Biol. 36, 52–64.

McCrea, H. J., and De Camilli, P. (2009). Mutations in phosphoinositide metabolizing enzymes and human disease. Physiol. Bethesda Md 24, 8–16.

McLaughlin, S., and Murray, D. (2005). Plasma membrane phosphoinositide organization by protein electrostatics. Nature 438, 605–611.

Mehta, Z. B., Pietka, G., and Lowe, M. (2014). The cellular and physiological functions of the Lowe syndrome protein OCRL1. Traffic Cph. Den. 15, 471–487.

Mellors, A., Lun, A. K., and Peled, O. N. (1975). Evidence for NAD nucleosidase in rabbit-liver lysosomes. Can. J. Biochem. 53, 143–148.

Mesmin, B., Bigay, J., Moser von Filseck, J., Lacas-Gervais, S., Drin, G., and Antonny, B. (2013). A Four-Step Cycle Driven by PI(4)P Hydrolysis Directs Sterol/PI(4)P Exchange by the ER-Golgi Tether OSBP. Cell 155, 830–843.

Miki, H., Miura, K., and Takenawa, T. (1996). N-WASP, a novel actin-depolymerizing protein, regulates the cortical cytoskeletal rearrangement in a PIP2-dependent manner downstream of tyrosine kinases. EMBO J. 15, 5326–5335.

Minogue, S., Chu, K. M. E., Westover, E. J., Covey, D. F., Hsuan, J. J., and Waugh, M. G. (2010). Relationship between phosphatidylinositol 4-phosphate synthesis, membrane organization, and lateral diffusion of PI4KIIalpha at the trans-Golgi network. J. Lipid Res. 51, 2314–2324.

Misawa, H., Ohtsubo, M., Copeland, N. G., Gilbert, D. J., Jenkins, N. A., and Yoshimura, A. (1998). Cloning and characterization of a novel class II phosphoinositide 3-kinase containing C2 domain. Biochem. Biophys. Res. Commun. 244, 531–539.

Mishra, A., Eathiraj, S., Corvera, S., and Lambright, D. G. (2010). Structural basis for Rab GTPase recognition and endosome tethering by the C2H2 zinc finger of Early Endosomal Autoantigen 1 (EEA1). Proc. Natl. Acad. Sci. U. S. A. 107, 10866–10871.

Mócsai, A., Ruland, J., and Tybulewicz, V. L. J. (2010). The SYK tyrosine kinase: a crucial player in diverse biological functions. Nat. Rev. Immunol. 10, 387–402.

Mondal, S., Ghosh-Roy, S., Loison, F., Li, Y., Jia, Y., Harris, C., Williams, D. A., and Luo, H. R. (2011). PTEN negatively regulates engulfment of apoptotic cells by modulating activation of Rac GTPase. J. Immunol. Baltim. Md 1950 187, 5783–5794.

Mooren, O. L., Galletta, B. J., and Cooper, J. A. (2012). Roles for actin assembly in endocytosis. Annu. Rev. Biochem. 81, 661–686.

Moser von Filseck, J., opi, A., Delfosse, V., Vanni, S., Jackson, C. L., Bourguet, W., and Drin, G. (2015). Phosphatidylserine transport by ORP/Osh proteins is driven by phosphatidylinositol 4-phosphate. Science 349, 432–436.

Mukae, N., Yokoyama, H., Yokokura, T., Sakoyama, Y., and Nagata, S. (2002). Activation of the innate immunity in Drosophila by endogenous chromosomal DNA that escaped apoptotic degradation. Genes Dev. 16, 2662–2671.

171

Mullins, R. D., Heuser, J. A., and Pollard, T. D. (1998). The interaction of Arp2/3 complex with actin: nucleation, high affinity pointed end capping, and formation of branching networks of filaments. Proc. Natl. Acad. Sci. U. S. A. 95, 6181–6186.

Murray, J. T., Panaretou, C., Stenmark, H., Miaczynska, M., and Backer, J. M. (2002). Role of Rab5 in the recruitment of hVps34/p150 to the early endosome. Traffic Cph. Den. 3, 416–427.

Nair-Gupta, P. et al. (2014). TLR signals induce phagosomal MHC-I delivery from the endosomal recycling compartment to allow cross-presentation. Cell 158, 506–521.

Nakatsu, F. et al. (2012). PtdIns4P synthesis by PI4KIIIα at the plasma membrane and its impact on plasma membrane identity. J. Cell Biol. 199, 1003–1016.

Nakatsu, F., Messa, M., Nández, R., Czapla, H., Zou, Y., Strittmatter, S. M., and De Camilli, P. (2015). Sac2/INPP5F is an inositol 4-phosphatase that functions in the endocytic pathway. J. Cell Biol. 209, 85–95.

Niedergang, F., Colucci-Guyon, E., Dubois, T., Raposo, G., and Chavrier, P. (2003). ADP ribosylation factor 6 is activated and controls membrane delivery during phagocytosis in macrophages. J. Cell Biol. 161, 1143–1150.

Niu, Y., Zhang, C., Sun, Z., Hong, Z., Li, K., Sun, D., Yang, Y., Tian, C., Gong, W., and Liu, J.-J. (2013). PtdIns(4)P regulates retromer–motor interaction to facilitate dynein–cargo dissociation at the trans-Golgi network. Nat. Cell Biol. 15, 417–429.

Nordmann, M., Cabrera, M., Perz, A., Bröcker, C., Ostrowicz, C., Engelbrecht-Vandré, S., and Ungermann, C. (2010). The Mon1-Ccz1 complex is the GEF of the late endosomal Rab7 homolog Ypt7. Curr. Biol. CB 20, 1654–1659.

Novakowski, K. E. et al. (2016). A naturally occurring transcript variant of MARCO reveals the SRCR domain is critical for function. Immunol. Cell Biol. 94, 646–655.

Nunes, P., Cornut, D., Bochet, V., Hasler, U., Oh-Hora, M., Waldburger, J.-M., and Demaurex, N. (2012). STIM1 juxtaposes ER to phagosomes, generating Ca2+ hotspots that boost phagocytosis. Curr. Biol. CB 22, 1990–1997.

Nunes, P., Demaurex, N., and Dinauer, M. C. (2013). Regulation of the NADPH oxidase and associated ion fluxes during phagocytosis. Traffic Cph. Den. 14, 1118–1131.

Oancea, E., Teruel, M. N., Quest, A. F., and Meyer, T. (1998). Green fluorescent protein (GFP)-tagged cysteine-rich domains from protein kinase C as fluorescent indicators for diacylglycerol signaling in living cells. J. Cell Biol. 140, 485–498.

Oikawa, T., Yamaguchi, H., Itoh, T., Kato, M., Ijuin, T., Yamazaki, D., Suetsugu, S., and Takenawa, T. (2004). PtdIns(3,4,5)P3 binding is necessary for WAVE2-induced formation of lamellipodia. Nat. Cell Biol. 6, 420–426.

Olmos, Y., and Carlton, J. G. (2016). The ESCRT machinery: new roles at new holes. Curr. Opin. Cell Biol. 38, 1–11.

Ostrowski, P. P., Fairn, G. D., Grinstein, S., and Johnson, D. E. (2016). Cresyl violet: a superior fluorescent lysosomal marker. Traffic Cph. Den.

172

Otsuka, A., Abe, T., Watanabe, M., Yagisawa, H., Takei, K., and Yamada, H. (2009). Dynamin 2 is required for actin assembly in phagocytosis in Sertoli cells. Biochem. Biophys. Res. Commun. 378, 478–482.

Padh, H., Ha, J., Lavasa, M., and Steck, T. L. (1993). A post-lysosomal compartment in Dictyostelium discoideum. J. Biol. Chem. 268, 6742–6747.

Palmieri, M., Nowell, C. J., Condron, M., Gardiner, J., Holmes, A. B., Desai, J., Burgess, A. W., and Catimel, B. (2010). Analysis of cellular phosphatidylinositol (3,4,5)-trisphosphate levels and distribution using confocal fluorescent microscopy. Anal. Biochem. 406, 41–50.

Park, H., and Cox, D. (2009). Cdc42 regulates Fc gamma receptor-mediated phagocytosis through the activation and phosphorylation of Wiskott-Aldrich syndrome protein (WASP) and neural-WASP. Mol. Biol. Cell 20, 4500–4508.

Patel, P. C., and Harrison, R. E. (2008). Membrane ruffles capture C3bi-opsonized particles in activated macrophages. Mol. Biol. Cell 19, 4628–4639.

Peplowska, K., Markgraf, D. F., Ostrowicz, C. W., Bange, G., and Ungermann, C. (2007). The CORVET tethering complex interacts with the yeast Rab5 homolog Vps21 and is involved in endo-lysosomal biogenesis. Dev. Cell 12, 739–750.

Pethe, K., Swenson, D. L., Alonso, S., Anderson, J., Wang, C., and Russell, D. G. (2004). Isolation of Mycobacterium tuberculosis mutants defective in the arrest of phagosome maturation. Proc. Natl. Acad. Sci. U. S. A. 101, 13642–13647.

Plemel, R. L., Lobingier, B. T., Brett, C. L., Angers, C. G., Nickerson, D. P., Paulsel, A., Sprague, D., and Merz, A. J. (2011). Subunit organization and Rab interactions of Vps-C protein complexes that control endolysosomal membrane traffic. Mol. Biol. Cell 22, 1353–1363.

Poteryaev, D., Datta, S., Ackema, K., Zerial, M., and Spang, A. (2010). Identification of the switch in early-to-late endosome transition. Cell 141, 497–508.

Pricop, L., Redecha, P., Teillaud, J. L., Frey, J., Fridman, W. H., Sautès-Fridman, C., and Salmon, J. E. (2001). Differential modulation of stimulatory and inhibitory Fc gamma receptors on human monocytes by Th1 and Th2 cytokines. J. Immunol. Baltim. Md 1950 166, 531–537.

Prod’Homme, V., Boyer, L., Dubois, N., Mallavialle, A., Munro, P., Mouska, X., Coste, I., Rottapel, R., Tartare-Deckert, S., and Deckert, M. (2015). Cherubism allele heterozygosity amplifies microbe-induced inflammatory responses in murine macrophages. J. Clin. Invest. 125, 1396–1400.

Pulecio, J., Tagliani, E., Scholer, A., Prete, F., Fetler, L., Burrone, O. R., and Benvenuti, F. (2008). Expression of Wiskott-Aldrich syndrome protein in dendritic cells regulates synapse formation and activation of naive CD8+ T cells. J. Immunol. Baltim. Md 1950 181, 1135–1142.

Puri, R. V., Reddy, P. V., and Tyagi, A. K. (2013). Secreted acid phosphatase (SapM) of Mycobacterium tuberculosis is indispensable for arresting phagosomal maturation and growth of the pathogen in guinea pig tissues. PloS One 8, e70514.

Rai, A., Pathak, D., Thakur, S., Singh, S., Dubey, A. K., and Mallik, R. (2016). Dynein Clusters into Lipid Microdomains on Phagosomes to Drive Rapid Transport toward Lysosomes. Cell 164, 722–734.

173

Rameh, L. E., and Cantley, L. C. (1999). The role of phosphoinositide 3-kinase lipid products in cell function. J. Biol. Chem. 274, 8347–8350.

Rebsamen, M. et al. (2015). SLC38A9 is a component of the lysosomal amino acid sensing machinery that controls mTORC1. Nature 519, 477–481.

Roberts, E. A., Chua, J., Kyei, G. B., and Deretic, V. (2006). Higher order Rab programming in phagolysosome biogenesis. J. Cell Biol. 174, 923–929.

Roberts, R. L., Barbieri, M. A., Ullrich, J., and Stahl, P. D. (2000). Dynamics of rab5 activation in endocytosis and phagocytosis. J. Leukoc. Biol. 68, 627–632.

Rocha, N., Kuijl, C., van der Kant, R., Janssen, L., Houben, D., Janssen, H., Zwart, W., and Neefjes, J. (2009). Cholesterol sensor ORP1L contacts the ER protein VAP to control Rab7-RILP-p150 Glued and late endosome positioning. J. Cell Biol. 185, 1209–1225.

Rodal, A. A., Sokolova, O., Robins, D. B., Daugherty, K. M., Hippenmeyer, S., Riezman, H., Grigorieff, N., and Goode, B. L. (2005). Conformational changes in the Arp2/3 complex leading to actin nucleation. Nat. Struct. Mol. Biol. 12, 26–31.

Roepstorff, K., Rasmussen, I., Sawada, M., Cudre-Maroux, C., Salmon, P., Bokoch, G., van Deurs, B., and Vilhardt, F. (2008). Stimulus-dependent regulation of the phagocyte NADPH oxidase by a VAV1, Rac1, and PAK1 signaling axis. J. Biol. Chem. 283, 7983–7993.

Rogers, N. C. et al. (2005). Syk-dependent cytokine induction by Dectin-1 reveals a novel pattern recognition pathway for C type lectins. Immunity 22, 507–517.

Rohatgi, R., Ho, H. Y., and Kirschner, M. W. (2000a). Mechanism of N-WASP activation by CDC42 and phosphatidylinositol 4, 5-bisphosphate. J. Cell Biol. 150, 1299–1310.

Rohatgi, R., Ho, H. Y., and Kirschner, M. W. (2000b). Mechanism of N-WASP activation by CDC42 and phosphatidylinositol 4, 5-bisphosphate. J. Cell Biol. 150, 1299–1310.

Rohde, H. M., Cheong, F. Y., Konrad, G., Paiha, K., Mayinger, P., and Boehmelt, G. (2003). The human phosphatidylinositol phosphatase SAC1 interacts with the coatomer I complex. J. Biol. Chem. 278, 52689–52699.

Rojas, R., van Vlijmen, T., Mardones, G. A., Prabhu, Y., Rojas, A. L., Mohammed, S., Heck, A. J. R., Raposo, G., van der Sluijs, P., and Bonifacino, J. S. (2008). Regulation of retromer recruitment to endosomes by sequential action of Rab5 and Rab7. J. Cell Biol. 183, 513–526.

Rong, Y. et al. (2012). Clathrin and phosphatidylinositol-4,5-bisphosphate regulate autophagic lysosome reformation. Nat. Cell Biol. 14, 924–934.

Rosa-Ferreira, C., and Munro, S. (2011). Arl8 and SKIP act together to link lysosomes to kinesin-1. Dev. Cell 21, 1171–1178.

Rutherford, A. C., Traer, C., Wassmer, T., Pattni, K., Bujny, M. V., Carlton, J. G., Stenmark, H., and Cullen, P. J. (2006). The mammalian phosphatidylinositol 3-phosphate 5-kinase (PIKfyve) regulates endosome-to-TGN retrograde transport. J. Cell Sci. 119, 3944–3957.

Saarikangas, J., Zhao, H., and Lappalainen, P. (2010). Regulation of the actin cytoskeleton-plasma membrane interplay by phosphoinositides. Physiol. Rev. 90, 259–289.

174

de Saint-Jean, M., Delfosse, V., Douguet, D., Chicanne, G., Payrastre, B., Bourguet, W., Antonny, B., and Drin, G. (2011). Osh4p exchanges sterols for phosphatidylinositol 4-phosphate between lipid bilayers. J. Cell Biol. 195, 965–978.

Salazar, G., Craige, B., Wainer, B. H., Guo, J., De Camilli, P., and Faundez, V. (2005). Phosphatidylinositol-4-kinase type II alpha is a component of adaptor protein-3-derived vesicles. Mol. Biol. Cell 16, 3692–3704.

Samejima, K., and Earnshaw, W. C. (2005). Trashing the genome: the role of nucleases during apoptosis. Nat. Rev. Mol. Cell Biol. 6, 677–688.

Sánchez-Barrena, M. J., Vallis, Y., Clatworthy, M. R., Doherty, G. J., Veprintsev, D. B., Evans, P. R., and McMahon, H. T. (2012). Bin2 is a membrane sculpting N-BAR protein that influences leucocyte podosomes, motility and phagocytosis. PloS One 7, e52401.

Sanematsu, F., Nishikimi, A., Watanabe, M., Hongu, T., Tanaka, Y., Kanaho, Y., Côté, J.-F., and Fukui, Y. (2013). Phosphatidic acid-dependent recruitment and function of the Rac activator DOCK1 during dorsal ruffle formation. J. Biol. Chem. 288, 8092–8100.

Saric, A., Hipolito, V. E. B., Kay, J. G., Canton, J., Antonescu, C. N., and Botelho, R. J. (2016). mTOR controls lysosome tubulation and antigen presentation in macrophages and dendritic cells. Mol. Biol. Cell 27, 321–333.

Sason, H., Milgrom, M., Weiss, A. M., Melamed-Book, N., Balla, T., Grinstein, S., Backert, S., Rosenshine, I., and Aroeti, B. (2009). Enteropathogenic Escherichia coli subverts phosphatidylinositol 4,5-bisphosphate and phosphatidylinositol 3,4,5-trisphosphate upon epithelial cell infection. Mol. Biol. Cell 20, 544–555.

Schlam, D., Bohdanowicz, M., Chatgilialoglu, A., Chatilialoglu, A., Steinberg, B. E., Ueyama, T., Du, G., Grinstein, S., and Fairn, G. D. (2013). Diacylglycerol kinases terminate diacylglycerol signaling during the respiratory burst leading to heterogeneous phagosomal NADPH oxidase activation. J. Biol. Chem. 288, 23090–23104.

Schneider, B. E., Behrends, J., Hagens, K., Harmel, N., Shayman, J. A., and Schaible, U. E. (2014). Lysosomal phospholipase A2: a novel player in host immunity to Mycobacterium tuberculosis. Eur. J. Immunol. 44, 2394–2404.

Scott, C. C., Dobson, W., Botelho, R. J., Coady-Osberg, N., Chavrier, P., Knecht, D. A., Heath, C., Stahl, P., and Grinstein, S. (2005). Phosphatidylinositol-4,5-bisphosphate hydrolysis directs actin remodeling during phagocytosis. J. Cell Biol. 169, 139–149.

Segal, A. W., Geisow, M., Garcia, R., Harper, A., and Miller, R. (1981). The respiratory burst of phagocytic cells is associated with a rise in vacuolar pH. Nature 290, 406–409.

Sender, R., Fuchs, S., and Milo, R. (2016). Are We Really Vastly Outnumbered? Revisiting the Ratio of Bacterial to Host Cells in Humans. Cell 164, 337–340.

Sheetz, M. P. (2001). Cell control by membrane-cytoskeleton adhesion. Nat. Rev. Mol. Cell Biol. 2, 392–396.

Shindo, M., Irie, K., Masuda, A., Ohigashi, H., Shirai, Y., Miyasaka, K., and Saito, N. (2003). Synthesis and phorbol ester binding of the cysteine-rich domains of diacylglycerol kinase (DGK)

175

isozymes. DGKgamma and DGKbeta are new targets of tumor-promoting phorbol esters. J. Biol. Chem. 278, 18448–18454.

Sicart, A., Katan, M., Egea, G., and Sarri, E. (2015). PLCγ1 participates in protein transport and diacylglycerol production triggered by cargo arrival at the Golgi. Traffic Cph. Den. 16, 250–266.

Simonsen, A., Gaullier, J. M., D’Arrigo, A., and Stenmark, H. (1999). The Rab5 effector EEA1 interacts directly with syntaxin-6. J. Biol. Chem. 274, 28857–28860.

Simonsen, A., Lippé, R., Christoforidis, S., Gaullier, J. M., Brech, A., Callaghan, J., Toh, B. H., Murphy, C., Zerial, M., and Stenmark, H. (1998). EEA1 links PI(3)K function to Rab5 regulation of endosome fusion. Nature 394, 494–498.

Simonsen, A., Wurmser, A. E., Emr, S. D., and Stenmark, H. (2001). The role of phosphoinositides in membrane transport. Curr. Opin. Cell Biol. 13, 485–492.

Sohn, M. et al. (2018). PI(4,5)P2 controls plasma membrane PI4P and PS levels via ORP5/8 recruitment to ER-PM contact sites. J. Cell Biol. 217, 1797–1813.

Stack, J. H., and Emr, S. D. (1994). Vps34p required for yeast vacuolar protein sorting is a multiple specificity kinase that exhibits both protein kinase and phosphatidylinositol-specific PI 3-kinase activities. J. Biol. Chem. 269, 31552–31562.

Stauffer, T. P., Ahn, S., and Meyer, T. (1998). Receptor-induced transient reduction in plasma membrane PtdIns(4,5)P2 concentration monitored in living cells. Curr. Biol. CB 8, 343–346.

Stein, M.-P., Feng, Y., Cooper, K. L., Welford, A. M., and Wandinger-Ness, A. (2003). Human VPS34 and p150 are Rab7 interacting partners. Traffic Cph. Den. 4, 754–771.

Stephens, L., Cooke, F. T., Walters, R., Jackson, T., Volinia, S., Gout, I., Waterfield, M. D., and Hawkins, P. T. (1994). Characterization of a phosphatidylinositol-specific phosphoinositide 3-kinase from mammalian cells. Curr. Biol. CB 4, 203–214.

Stuart, L. M., and Ezekowitz, R. A. B. (2005). Phagocytosis. Immunity 22, 539–550.

Suchanek, M., Hynynen, R., Wohlfahrt, G., Lehto, M., Johansson, M., Saarinen, H., Radzikowska, A., Thiele, C., and Olkkonen, V. M. (2007). The mammalian oxysterol-binding protein-related proteins (ORPs) bind 25-hydroxycholesterol in an evolutionarily conserved pocket. Biochem. J. 405, 473–480.

Suetsugu, S., Kurisu, S., Oikawa, T., Yamazaki, D., Oda, A., and Takenawa, T. (2006). Optimization of WAVE2 complex-induced actin polymerization by membrane-bound IRSp53, PIP(3), and Rac. J. Cell Biol. 173, 571–585.

Suh, B.-C., and Hille, B. (2005). Regulation of ion channels by phosphatidylinositol 4,5-bisphosphate. Curr. Opin. Neurobiol. 15, 370–378.

Swanson, J. A. (2004). The coordination of signaling during Fc receptor-mediated phagocytosis. J. Leukoc. Biol. 76, 1093–1103.

Swanson, J. A. (2008). Shaping cups into phagosomes and macropinosomes. Nat. Rev. Mol. Cell Biol. 9, 639–649.

176

Swanson, J. A. (2014). Phosphoinositides and engulfment. Cell. Microbiol. 16, 1473–1483.

Swanson, J. A., Johnson, M. T., Beningo, K., Post, P., Mooseker, M., and Araki, N. (1999). A contractile activity that closes phagosomes in macrophages. J. Cell Sci. 112 ( Pt 3), 307–316.

Syam, S., Mero, P., Pham, T., McIntosh, C. A., Bruhns, P., and Booth, J. W. (2010). Differential recruitment of activating and inhibitory Fc gamma RII during phagocytosis. J. Immunol. Baltim. Md 1950 184, 2966–2973.

Szentpetery, Z., Várnai, P., and Balla, T. (2010). Acute manipulation of Golgi phosphoinositides to assess their importance in cellular trafficking and signaling. Proc. Natl. Acad. Sci. U. S. A. 107, 8225–8230.

Takano, K., Takano, K., Toyooka, K., and Suetsugu, S. (2008). EFC/F-BAR proteins and the N-WASP-WIP complex induce membrane curvature-dependent actin polymerization. EMBO J. 27, 2817–2828.

Tan, J., and Brill, J. A. (2014). Cinderella story: PI4P goes from precursor to key signaling molecule. Crit. Rev. Biochem. Mol. Biol. 49, 33–58.

Terebiznik, M. R., Vieira, O. V., Marcus, S. L., Slade, A., Yip, C. M., Trimble, W. S., Meyer, T., Finlay, B. B., and Grinstein, S. (2002). Elimination of host cell PtdIns(4,5)P(2) by bacterial SigD promotes membrane fission during invasion by Salmonella. Nat. Cell Biol. 4, 766–773.

Tian, W., Li, X. J., Stull, N. D., Ming, W., Suh, C.-I., Bissonnette, S. A., Yaffe, M. B., Grinstein, S., Atkinson, S. J., and Dinauer, M. C. (2008). Fc gamma R-stimulated activation of the NADPH oxidase: phosphoinositide-binding protein p40phox regulates NADPH oxidase activity after enzyme assembly on the phagosome. Blood 112, 3867–3877.

Tolias, K. F., Cantley, L. C., and Carpenter, C. L. (1995). Rho family GTPases bind to phosphoinositide kinases. J. Biol. Chem. 270, 17656–17659.

Tolias, K. F., Hartwig, J. H., Ishihara, H., Shibasaki, Y., Cantley, L. C., and Carpenter, C. L. (2000). Type Ialpha phosphatidylinositol-4-phosphate 5-kinase mediates Rac-dependent actin assembly. Curr. Biol. CB 10, 153–156.

Tolias, K. F., Rameh, L. E., Ishihara, H., Shibasaki, Y., Chen, J., Prestwich, G. D., Cantley, L. C., and Carpenter, C. L. (1998). Type I phosphatidylinositol-4-phosphate 5-kinases synthesize the novel lipids phosphatidylinositol 3,5-bisphosphate and phosphatidylinositol 5-phosphate. J. Biol. Chem. 273, 18040–18046.

Tomasevic, N., Jia, Z., Russell, A., Fujii, T., Hartman, J. J., Clancy, S., Wang, M., Beraud, C., Wood, K. W., and Sakowicz, R. (2007). Differential regulation of WASP and N-WASP by Cdc42, Rac1, Nck, and PI(4,5)P2. Biochemistry (Mosc.) 46, 3494–3502.

Tong, J., Yang, H., Yang, H., Eom, S. H., and Im, Y. J. (2013). Structure of Osh3 reveals a conserved mode of phosphoinositide binding in oxysterol-binding proteins. Struct. Lond. Engl. 1993 21, 1203–1213.

Trajkovic, K., Hsu, C., Chiantia, S., Rajendran, L., Wenzel, D., Wieland, F., Schwille, P., Brügger, B., and Simons, M. (2008). Ceramide triggers budding of exosome vesicles into multivesicular endosomes. Science 319, 1244–1247.

177

Treanor, B., Depoil, D., Gonzalez-Granja, A., Barral, P., Weber, M., Dushek, O., Bruckbauer, A., and Batista, F. D. (2010). The membrane skeleton controls diffusion dynamics and signaling through the B cell receptor. Immunity 32, 187–199.

Tridandapani, S., Lyden, T. W., Smith, J. L., Carter, J. E., Coggeshall, K. M., and Anderson, C. L. (2000). The adapter protein LAT enhances fcgamma receptor-mediated signal transduction in myeloid cells. J. Biol. Chem. 275, 20480–20487.

Trimble, W. S., and Grinstein, S. (2015). Barriers to the free diffusion of proteins and lipids in the plasma membrane. J. Cell Biol. 208, 259–271.

Trost, M., English, L., Lemieux, S., Courcelles, M., Desjardins, M., and Thibault, P. (2009). The phagosomal proteome in interferon-gamma-activated macrophages. Immunity 30, 143–154.

Truman, L. A. et al. (2008). CX3CL1/fractalkine is released from apoptotic lymphocytes to stimulate macrophage chemotaxis. Blood 112, 5026–5036.

Tse, S. M. L., Furuya, W., Gold, E., Schreiber, A. D., Sandvig, K., Inman, R. D., and Grinstein, S. (2003). Differential role of actin, clathrin, and dynamin in Fc gamma receptor-mediated endocytosis and phagocytosis. J. Biol. Chem. 278, 3331–3338.

Tsuboi, S. et al. (2009). FBP17 Mediates a Common Molecular Step in the Formation of Podosomes and Phagocytic Cups in Macrophages. J. Biol. Chem. 284, 8548–8556.

Tsujita, K., Itoh, T., Ijuin, T., Yamamoto, A., Shisheva, A., Laporte, J., and Takenawa, T. (2004). Myotubularin regulates the function of the late endosome through the gram domain-phosphatidylinositol 3,5-bisphosphate interaction. J. Biol. Chem. 279, 13817–13824.

Tsujita, K., Suetsugu, S., Sasaki, N., Furutani, M., Oikawa, T., and Takenawa, T. (2006). Coordination between the actin cytoskeleton and membrane deformation by a novel membrane tubulation domain of PCH proteins is involved in endocytosis. J. Cell Biol. 172, 269–279.

Uderhardt, S. et al. (2012). 12/15-lipoxygenase orchestrates the clearance of apoptotic cells and maintains immunologic tolerance. Immunity 36, 834–846.

Ueyama, T. et al. (2004). Superoxide production at phagosomal cup/phagosome through beta I protein kinase C during Fc gamma R-mediated phagocytosis in microglia. J. Immunol. Baltim. Md 1950 173, 4582–4589.

Ueyama, T., Kusakabe, T., Karasawa, S., Kawasaki, T., Shimizu, A., Son, J., Leto, T. L., Miyawaki, A., and Saito, N. (2008). Sequential binding of cytosolic Phox complex to phagosomes through regulated adaptor proteins: evaluation using the novel monomeric Kusabira-Green System and live imaging of phagocytosis. J. Immunol. Baltim. Md 1950 181, 629–640.

Ueyama, T., Tatsuno, T., Kawasaki, T., Tsujibe, S., Shirai, Y., Sumimoto, H., Leto, T. L., and Saito, N. (2007). A regulated adaptor function of p40phox: distinct p67phox membrane targeting by p40phox and by p47phox. Mol. Biol. Cell 18, 441–454.

Underhill, D. M. (2005). Phagosome maturation: steady as she goes. Immunity 23, 343–344.

Underhill, D. M., and Gantner, B. (2004). Integration of Toll-like receptor and phagocytic signaling for tailored immunity. Microbes Infect. 6, 1368–1373.

178

Underhill, D. M., and Goodridge, H. S. (2012). Information processing during phagocytosis. Nat. Rev. Immunol. 12, 492–502.

Underhill, D. M., and Pearlman, E. (2015). Immune Interactions with Pathogenic and Commensal Fungi: A Two-Way Street. Immunity 43, 845–858.

Underhill, D. M., Rossnagle, E., Lowell, C. A., and Simmons, R. M. (2005). Dectin-1 activates Syk tyrosine kinase in a dynamic subset of macrophages for reactive oxygen production. Blood 106, 2543–2550.

Utomo, A., Cullere, X., Glogauer, M., Swat, W., and Mayadas, T. N. (2006). Vav proteins in neutrophils are required for FcgammaR-mediated signaling to Rac GTPases and nicotinamide adenine dinucleotide phosphate oxidase component p40(phox). J. Immunol. Baltim. Md 1950 177, 6388–6397.

Vanhaesebroeck, B., and Waterfield, M. D. (1999). Signaling by distinct classes of phosphoinositide 3-kinases. Exp. Cell Res. 253, 239–254.

Vargas, P. et al. (2016). Innate control of actin nucleation determines two distinct migration behaviours in dendritic cells. Nat. Cell Biol. 18, 43–53.

Vergne, I., Chua, J., and Deretic, V. (2003). Tuberculosis toxin blocking phagosome maturation inhibits a novel Ca2+/calmodulin-PI3K hVPS34 cascade. J. Exp. Med. 198, 653–659.

Vieira, O. V., Botelho, R. J., Rameh, L., Brachmann, S. M., Matsuo, T., Davidson, H. W., Schreiber, A., Backer, J. M., Cantley, L. C., and Grinstein, S. (2001). Distinct roles of class I and class III phosphatidylinositol 3-kinases in phagosome formation and maturation. J. Cell Biol. 155, 19–25.

Vieira, O. V., Bucci, C., Harrison, R. E., Trimble, W. S., Lanzetti, L., Gruenberg, J., Schreiber, A. D., Stahl, P. D., and Grinstein, S. (2003). Modulation of Rab5 and Rab7 recruitment to phagosomes by phosphatidylinositol 3-kinase. Mol. Cell. Biol. 23, 2501–2514.

Vieira, O. V., Harrison, R. E., Scott, C. C., Stenmark, H., Alexander, D., Liu, J., Gruenberg, J., Schreiber, A. D., and Grinstein, S. (2004). Acquisition of Hrs, an essential component of phagosomal maturation, is impaired by mycobacteria. Mol. Cell. Biol. 24, 4593–4604.

Vihervaara, T., Uronen, R.-L., Wohlfahrt, G., Björkhem, I., Ikonen, E., and Olkkonen, V. M. (2011). Sterol binding by OSBP-related protein 1L regulates late endosome motility and function. Cell. Mol. Life Sci. CMLS 68, 537–551.

Voronov, S. V. et al. (2008). Synaptojanin 1-linked phosphoinositide dyshomeostasis and cognitive deficits in mouse models of Down’s syndrome. Proc. Natl. Acad. Sci. U. S. A. 105, 9415–9420.

Wähe, A., Kasmapour, B., Schmaderer, C., Liebl, D., Sandhoff, K., Nykjaer, A., Griffiths, G., and Gutierrez, M. G. (2010). Golgi-to-phagosome transport of acid sphingomyelinase and prosaposin is mediated by sortilin. J. Cell Sci. 123, 2502–2511.

Wang, H., Sun, H.-Q., Zhu, X., Zhang, L., Albanesi, J., Levine, B., and Yin, H. (2015a). GABARAPs regulate PI4P-dependent autophagosome:lysosome fusion. Proc. Natl. Acad. Sci. U. S. A. 112, 7015–7020.

179

Wang, S. et al. (2015b). Metabolism. Lysosomal amino acid transporter SLC38A9 signals arginine sufficiency to mTORC1. Science 347, 188–194.

Wang, Y. J., Wang, J., Sun, H. Q., Martinez, M., Sun, Y. X., Macia, E., Kirchhausen, T., Albanesi, J. P., Roth, M. G., and Yin, H. L. (2003). Phosphatidylinositol 4 phosphate regulates targeting of clathrin adaptor AP-1 complexes to the Golgi. Cell 114, 299–310.

Weernink, P. A. O., Meletiadis, K., Hommeltenberg, S., Hinz, M., Ishihara, H., Schmidt, M., and Jakobs, K. H. (2004). Activation of type I phosphatidylinositol 4-phosphate 5-kinase isoforms by the Rho GTPases, RhoA, Rac1, and Cdc42. J. Biol. Chem. 279, 7840–7849.

Welch, M. D., Rosenblatt, J., Skoble, J., Portnoy, D. A., and Mitchison, T. J. (1998). Interaction of human Arp2/3 complex and the Listeria monocytogenes ActA protein in actin filament nucleation. Science 281, 105–108.

Whitters, E. A., Cleves, A. E., McGee, T. P., Skinner, H. B., and Bankaitis, V. A. (1993). SAC1p is an integral membrane protein that influences the cellular requirement for phospholipid transfer protein function and inositol in yeast. J. Cell Biol. 122, 79–94.

Wijdeven, R. H., Janssen, H., Nahidiazar, L., Janssen, L., Jalink, K., Berlin, I., and Neefjes, J. (2016). Cholesterol and ORP1L-mediated ER contact sites control autophagosome transport and fusion with the endocytic pathway. Nat. Commun. 7, 11808.

Wolfson, R. L., Chantranupong, L., Saxton, R. A., Shen, K., Scaria, S. M., Cantor, J. R., and Sabatini, D. M. (2016). Sestrin2 is a leucine sensor for the mTORC1 pathway. Science 351, 43–48.

Wollert, T., and Hurley, J. H. (2010). Molecular mechanism of multivesicular body biogenesis by ESCRT complexes. Nature 464, 864–869.

Wong, H. S., Jaumouillé, V., Freeman, S. A., Doodnauth, S. A., Schlam, D., Canton, J., Mukovozov, I. M., Saric, A., Grinstein, S., and Robinson, L. A. (2016). Chemokine Signaling Enhances CD36 Responsiveness toward Oxidized Low-Density Lipoproteins and Accelerates Foam Cell Formation. Cell Rep.

Wu, X., Chi, R. J., Baskin, J. M., Lucast, L., Burd, C. G., De Camilli, P., and Reinisch, K. M. (2014). Structural insights into assembly and regulation of the plasma membrane phosphatidylinositol 4-kinase complex. Dev. Cell 28, 19–29.

Yamada, H. et al. (2007). Amphiphysin 1 is important for actin polymerization during phagocytosis. Mol. Biol. Cell 18, 4669–4680.

Yamauchi, S., Kawauchi, K., and Sawada, Y. (2012). Myosin II-dependent exclusion of CD45 from the site of Fcγ receptor activation during phagocytosis. FEBS Lett. 586, 3229–3235.

Yan, Y., and Backer, J. M. (2007). Regulation of class III (Vps34) PI3Ks. Biochem. Soc. Trans. 35, 239–241.

Yeung, T., Terebiznik, M., Yu, L., Silvius, J., Abidi, W. M., Philips, M., Levine, T., Kapus, A., and Grinstein, S. (2006). Receptor activation alters inner surface potential during phagocytosis. Science 313, 347–351.

180

Yoshida, S., Pacitto, R., Yao, Y., Inoki, K., and Swanson, J. A. (2015). Growth factor signaling to mTORC1 by amino acid-laden macropinosomes. J. Cell Biol. 211, 159–172.

Yu, L. et al. (2010). Termination of autophagy and reformation of lysosomes regulated by mTOR. Nature 465, 942–946.

Yu, M., Lowell, C. A., Neel, B. G., and Gu, H. (2006). Scaffolding adapter Grb2-associated binder 2 requires Syk to transmit signals from FcepsilonRI. J. Immunol. Baltim. Md 1950 176, 2421–2429.

Zencheck, W. D., Xiao, H., Nolen, B. J., Angeletti, R. H., Pollard, T. D., and Almo, S. C. (2009). Nucleotide- and activator-dependent structural and dynamic changes of arp2/3 complex monitored by hydrogen/deuterium exchange and mass spectrometry. J. Mol. Biol. 390, 414–427.

Zewe, J. P., Wills, R. C., Sangappa, S., Goulden, B. D., and Hammond, G. R. (2018). SAC1 degrades its lipid substrate PtdIns4P in the endoplasmic reticulum to maintain a steep chemical gradient with donor membranes. ELife 7.

Zhang, L., Malik, S., Pang, J., Wang, H., Park, K. M., Yule, D. I., Blaxall, B. C., and Smrcka, A. V. (2013). Phospholipase Cε hydrolyzes perinuclear phosphatidylinositol 4-phosphate to regulate cardiac hypertrophy. Cell 153, 216–227.

Zhang, Z., Zhang, T., Wang, S., Gong, Z., Tang, C., Chen, J., and Ding, J. (2014). Molecular mechanism for Rabex-5 GEF activation by Rabaptin-5. ELife 3.

Zhao, K., and Ridgway, N. D. (2017). Oxysterol-Binding Protein-Related Protein 1L Regulates Cholesterol Egress from the Endo-Lysosomal System. Cell Rep. 19, 1807–1818.

Zhu, H., Liang, Z., and Li, G. (2009). Rabex-5 is a Rab22 effector and mediates a Rab22-Rab5 signaling cascade in endocytosis. Mol. Biol. Cell 20, 4720–4729.

Zizzo, G., Hilliard, B. A., Monestier, M., and Cohen, P. L. (2012). Efficient clearance of early apoptotic cells by human macrophages requires M2c polarization and MerTK induction. J. Immunol. Baltim. Md 1950 189, 3508–3520.

Zoncu, R., Bar-Peled, L., Efeyan, A., Wang, S., Sancak, Y., and Sabatini, D. M. (2011). mTORC1 senses lysosomal amino acids through an inside-out mechanism that requires the vacuolar H(+)-ATPase. Science 334, 678–683.

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Appendix I

Appendix I. Phosphoinositide metabolism in mammalian cells. Diagram showing the metabolic interconversion of phosphoinositides in mammalian cells. Structures of individual phosphoinositides are shown with their respective nomenclatures in italics. Lipid kinases are shown in blue and lipid phosphatases are shown in red. Dotted lines represent reactions that have not been characterized in cells.

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Copyright Acknowledgements

Chapter 1 includes material published in “Biochimica et Biophysica Acta – Molecular and Cell

Biology of Lipids”. As an author of the article, the right to include it in this dissertation is retained.

Roni Levin, Sergio Grinstein and Daniel Schlam. “Phosphoinositides in Phagocytosis and

Macropinocytosis.” Biochimica et Biophysica Acta – Molecular and Cell Biology of Lipids.

September 2014.

Chapter 1 includes material published in “Immunological Reviews”. A license agreement

(#4427081282509) has been obtained to reuse the contents of this article.

Roni Levin, Sergio Grinstein and Johnathan Canton. “The life cycle of phagosomes: formation,

maturation and resolution.” Immunological Reviews – Neutrophils. August 2016.

Chapter 3 includes material published in “Springer Nature – Methods in Molecular Biology”. A

license agreement (#4427091035795) has been obtained to reuse the contents of this article.

Fernando Montaño, Sergio Grinstein and Roni Levin. ”Quantitative phagocytosis assays in

primary and cultures macrophages.” Methods in Molecular Biology, 2018;1784:151-163.

Chapter 3 includes material originally published in “Molecular Biology of the Cell”. A license

agreement (#4427260112162) has been obtained to reuse the contents of this article.

Roni Levin, Gerald R.V. Hammond, Tamas Balla, Pietro de Camilli, Gregory, D. Fairn and

Sergio Grinstein. “Multiphasic dynamics of phosphatidylinositol 4-phosphate during

phagocytosis.” Molecular Biology of the Cell, 2017, Jan 1;28(1):128-140.

Chapter 4 includes material originally published in “Molecular Biology of the Cell”. A license

agreement (#4427260112162) has been obtained to reuse the contents of this article.

Roni Levin, Gerald R.V. Hammond, Tamas Balla, Pietro de Camilli, Gregory, D. Fairn and

Sergio Grinstein. “Multiphasic dynamics of phosphatidylinositol 4-phosphate during

phagocytosis.” Molecular Biology of the Cell, 2017, Jan 1;28(1):128-140.