Investigating the Functional Roles of NBEAL2 in Megakaryocytes and Platelets

Investigating the functional roles of NBEAL2 in megakaryocytes and platelets

Richard Wei-Chi Lo

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

Department of Biochemistry University of Toronto

Investigating the functional roles of NBEAL2 in megakaryocytes and platelets

Richard Wei-Chi Lo

Doctor of Philosophy

Department of Biochemistry

University of Toronto

2019

Abstract

Platelets are small anucleate blood cells that co-ordinate blood clotting in response to vascular injury. In additional to their hemostatic role, platelets are involved in a diverse range of physiological and pathological processes. Platelets contain abundant secretory vesicles associated with their functions, which are generally classed as α-granules, δ-granules and lysosomes. These vesicles originate in megakaryocytes (MK), hematopoietic cells that produce platelets in the bone marrow. The inherited bleeding disorder Gray Platelet

Syndrome (GPS) is characterized by platelet α-granule deficiency arising from loss of function of neurobeachin-like 2 (NBEAL2). The goal of my study was to investigate the roles and functions of NBEAL2 in MKs and platelets.

Using biochemical assays, it was found that NBEAL2 is a cytosolic protein capable of self-association and adopts two distinct oligomeric states in MKs. Immunoprecipitation-mass spectrometry was used to identify several potential NBEAL2-interacting proteins, providing insights into its cellular functions. Among these interactors, SEC22B was shown to be essential for the formation of α-granules, where loss of SEC22B expression resulted in an α- granule deficiency similar to that caused by the loss of NBEAL2. Forms of NBEAL2 expressing the GPS-associated mutations E1833K or R1839C failed to interact with SEC22B, indicating a functional significance for NBEAL2-SEC22B interactions in α-granule biogenesis.

A Nbeal2-/- mouse strain was generated to model GPS. Studies of Nbeal2-/- mice revealed that NBEAL2 is required for normal platelet activation and aggregation, and for normal development and proplatelet-platelet production by MKs. Studies of the uptake and intracellular trafficking of the α-granule cargo protein fibrinogen (FGN) revealed that NBEAL2 is not essential for FGN endocytosis by MKs, nor its passage through endocytic compartments associated with RAB5, RAB7 and the α-granule membrane protein P-selectin.

The latter compartment retains both endocytosed and MK-synthesized cargo proteins when

NBEAL2 is present, but in cells lacking NBEAL2 cargo proteins are lost from this compartment and exit MKs via a RAB11-mediated secretory pathway. Intracellular protein colocalization and co-immunoprecipitation studies revealed a potential interaction between

NBEAL2 and P-selectin in MKs and platelets. Thus we propose that NBEAL2 is required for the stabilization of α-granules during cargo loading, maturation and circulation within platelets.

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Acknowledgements

I would like to express my sincere and deepest gratitude to my mentor Dr. Walter Kahr for his guidance, enthusiasm, inspiration, motivation, and knowledge. Thank you for the continuous support throughout the years of my graduate experience. I would like to thank my committee member Dr. William Trimble for his insight, advice and support over the years of my graduate study. I am grateful to Dr. Peter Kim for serving in my committee and for his insightful discussions. I would like to extend my appreciation to all the members of my supervisory committee for advancing my personal and professional developments.

The work presented in this thesis was made possible only because of the contributions from the members of the Kahr lab. My experiences in the lab are incredible highlights of my graduate study. I am especially grateful to Dr. Ling Li and Dr. Pluthero for their technical expertise, helpful insights and constant assistances. I would also like to thank all members of the Trimble lab for their helpful inputs and sharing of techniques and reagents.

Table of Contents

Acknowledgments ...... iii Table of Contents ...... v List of Tables ...... xi List of Figures ...... xii List of Abbreviations ...... xv

Chapter 1 Introduction 1.1 Platelet structure and function 1.1.1 Overview ...... 2 1.1.2 Roles of platelets in hemostasis ...... 3 1.1.3 Other physiological roles of platelets ...... 6 1.1.4 Pathological roles of platelets 1.1.4.1 Platelets, atheroscelerosis and thrombosis ...... 8 1.1.4.2 Platelets and cancer ...... 13 1.2 Megakaryocytes and platelet biogenesis ...... 15 1.3 Platelet structures and secretory granules 1.3.1 Overview ...... 18 1.3.2 δ-granule biogenesis in megakaryocytes and platelets 1.3.2.1 Overview ...... 20 1.3.2.2 Platelet δ-granule deficiencies 1.3.2.2.1 Hermansky-Pudlak syndrome ...... 20 1.3.2.2.2 Chediak-Higashi syndrome ...... 23 1.3.2.2.3 Griscelli syndrome ...... 24 1.3.2.3 Model of δ-granule biogenesis ...... 26 1.3.3 α-granule biogenesis in megakaryocytes and platelets 1.3.3.1 Overview ...... 27 1.3.3.2 α-granule trafficking 1.3.3.2.1 Trafficking of α-granule proteins ...... 28 1.3.3.2.2 Endosome maturation ...... 29 1.3.3.3 Platelet α-granule deficiencies 1.3.3.3.1 Gray platelet syndrome ...... 31 1.3.3.3.2 Arthrogryposis, renal dysfunction and cholestasis syndrome

...... 35 1.3.3.4 Model of α-granule biogenesis ...... 36 1.4 NBEAL2: a protein with unknown functions 1.4.1 PH-BEACH protein family ...... 38 1.4.2 NBEAL2 in megakaryocytes ...... 41 1.5 Rationale, Hypothesis and Aim ...... 43

Chapter 2 Materials and Methods 2.1 Human megakaryocytes and platelets ...... 45 2.2 Knockout and transgenic mice ...... 45 2.3 Cell lines and cultures ...... 46 2.3.1 Isolation and culture of human primary CD34+ cells ...... 46 2.3.2 Isolation and culture of mouse primary bone marrow cells ...... 47 2.3.3 Purification of human and mouse primary megakaryocytes ...... 47 2.3.4 Isolation of human and mouse platelets ...... 48 2.4 Antibodies ...... 48 2.5 Constructs 2.5.1 Bacterial expression constructs ...... 49 2.5.2 Yeast expression constructs ...... 50 2.5.3 Mammalian expression constructs ...... 50 2.5.4 Lentiviral constructs ...... 51 2.6 Transfection and electroporation ...... 51 2.7 Lentivirus production and transduction 2.7.1 Protein expression in mammalian cell lines and human primary CD34+ cells ... 52 2.7.2 CRISPR knockout in mammalian cells ...... 53 2.8 Yeast two-hybrid ...... 54 2.9 Western blotting ...... 55 2.10 Immunoprecipitation ...... 55 2.11 Blue Native gel electrophoresis ...... 56 2.12 Cell fractionation ...... 56 2.13 Mass spectrometry and analysis ...... 57 2.14 Expression and purification of NBEAL2 PH-BEACH domain ...... 58 2.15 Fibrinogen endocytosis assays in mouse primary megakaryocytes ...... 59 2.16 Biotinylated fibrinogen for secretion and degradation assays ...... 59

2.17 Flow cytometry and analysis ...... 60 2.18 Imaging 2.18.1 Transmission electron and immunoelectron microscopy ...... 60 2.18.2 Confocal immunofluorescence microscopy ...... 61 2.19 Statistics ...... 62

Chapter 3 Investigating the molecular and structural properties of NBEAL2 3.1 Abstract ...... 64 3.2 Introduction ...... 64 3.3 Results 3.3.1 NBEAL2 forms oligomeric complexes ...... 64 3.3.2 Validation of NBEAL2 antibody ...... 66 3.3.3 NBEAL2 forms a single molecular weight complex in DAMI cells ...... 70 3.3.4 Two distinct NBEAL2 complexes in primary human megakaryocytes are both cytosolic and membrane-associated ...... 73 3.3.5 NBEAL2 localizes primarily to the α-granule marker, P-selectin, in human primary megakaryocytes ...... 75 3.3.6 NBEAL2 interacts with P-selectin ...... 77 3.3.7 Expression and purification of the PH-BEACH domain of NBEAL2 ...... 79 3.4 Discussion 3.4.1 NBEAL2 and PH-BEACH family proteins as oligomeric complexes ...... 86 3.4.2 NBEAL2 displays variable properties in DAMI cells and in primary human megakaryocytes ...... 87 3.4.3 Distinct configuration states of NBEAL2 in the cytosol and with membranes .... 88 3.4.4 NBEAL2 functions at α-granules ...... 89 3.4.5 PH-BEACH domain may direct the localization of NBEAL2 ...... 89 3.4.6 GPS mutations in the PH-BEACH domain provide mechanistic clues ...... 90

Chapter 4 Identification and characterization of the binding partners of NBEAL2 4.1 Abstract ...... 92 4.2 Introduction ...... 92 4.3 Results 4.3.1 Identification of NBEAL2 binding partners using yeast-two-hybrid screens ...... 92

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4.3.2 Identification of NBEAL2 binding partners in DAMI cells using affinity purification mass spectrometry ...... 96 4.3.3 Verification of NBEAL2 binding partners ...... 97 4.3.4 NBEAL2 and SEC22B co-localizes in primary human megakaryocytes ...... 100 4.3.5 Delineation of binding region of NBEAL2 to SEC22B ...... 102 4.3.6 NBEAL2 contains an inhibitory domain that suppresses NBEAL2-SEC22B interaction ...... 105 4.3.7 NBEAL2 protein level is dependent on the presence of SEC22B ...... 108 4.3.8 NBEAL2 and SEC22B are both required for α-granule biogenesis ...... 110 4.3.9 NBEAL2-SEC22B interaction is essential for α-granule formation ...... 111 4.4 Discussion 4.4.1 Binding partners provide insights into NBEAL2 function ...... 114 4.4.2 Dynamic interaction between NBEAL2 and SEC22B ...... 116 4.4.3 SEC22B affects NBEAL2 protein levels ...... 118 4.4.4 NBEAL2 and SEC22B are critical to α-granule biogenesis ...... 118

Chapter 5 NBEAL2 is required for normal development of megakaryocytes 5.1 Abstract ...... 121 5.2 Introduction ...... 121 5.3 Results 5.3.1 Nbeal2-/- mice as a model of human GPS ...... 122 5.3.2 Nbeal2-/- platelets contain reduced α-granule cargo and membrane proteins . 124 5.3.3 Structural and functional abnormalities of Nbeal2-/- megakaryocytes and platelets ...... 125 5.3.4 Absence of Nbeal2 leads to defects in megakaryocyte development and proplatelet formation ...... 130 5.4 Discussion 5.4.1 NBEAL2 knockout leads to phenotypes that resemble human GPS in mice ... 141 5.4.2 Nbeal2-/- platelets are functionally defective and contribute to bleeding in mice ...... 142 5.4.3 Nbeal2-/- platelets lack α-granule cargo proteins but retain membrane components of α-granules ...... 143 5.4.4 NBEAL2 may be required for the stabilization of α-granules ...... 143

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5.4.5 NBEAL2 is required for normal megakaryocyte development and platelet production ...... 144

Chapter 6 Role of NBEAL2 in α-granule cargo trafficking 6.1 Abstract ...... 147 6.2 Introduction ...... 147 6.3 Results 6.3.1 Aberrant α-granule cargo protein trafficking in Nbeal2-/- megakaryocytes ...... 147 6.3.2 Expression of fluorescent NBEAL2 in Nbeal2-/- mice ...... 150 6.3.3 Intracellular fibrinogen is significantly reduced in Nbeal2-/- megakaryocytes following endocytosis ...... 153 6.3.4 Endocytosis is not affected by the loss of NBEAL2 ...... 155 6.3.5 Fibrinogen is not lost as a result of lysosomal degradation in Nbeal2-/- megakaryocytes ...... 157 6.3.6 Absence of NBEAL2 results in aberrant secretion of fibrinogen ...... 158 6.3.7 Fibrinogen is not retained in P-selectin positive vesicles but is secreted via Rab11+ vesicles in the absence of NBEAL2 ...... 159 6.3.8 Endogenously synthesized α-granule protein, VWF, is aberrantly trafficked to Rab11+ vesicles in Nbeal2-/- megakaryocytes ...... 164 6.3.9 P-selectin was not detected on the surface of Nbeal2-/- megakaryocytes with VWF ...... 166 6.4 Discussion 6.4.1 Absence of NBEAL2 results in the aberrant loss of α-granule cargo proteins in megakaryocytes ...... 168 6.4.2 Rescue of Nbeal2-/- mice with fluorescent NBEAL2 ...... 168 6.4.3 Loss of fibrinogen in Nbeal2-/- megakaryocytes is not caused by aberrant endocytosis or as a result of enhanced degradation ...... 169 6.4.4 Lack of NBEAL2 results in aberrant secretion of fibrinogen ...... 169 6.4.5 Fibrinogen and VWF are abnormally trafficked in Nbeal2-/- megakaryocytes .. 169 6.4.6 α-granules externalization is not the direct cause of α-granule protein loss .... 170 6.4.7 NBEAL2 is required for the retention of α-granule proteins ...... 171

Chapter 7 Concluding remarks 7.1 Functional significance of NBEAL2 molecular and structural properties ...... 174

7.2 SEC22B is a novel interacting partner of NBEAL2 that is required for α-granule biogenesis...... 175 7.3 A model for the role of NBEAL2 in α-granule biogenesis ...... 175 7.4 Implications in megakaryocyte and platelet biology ...... 176 7.5 Future directions 7.5.1 Molecular and structural properties of NBEAL ...... 178 7.5.2 SEC22B as an essential binding partner of NBEAL2 in α-granule biogenesis 179 7.5.3 NBEAL2 stabilizes α-granules to retain α-granule cargo proteins ...... 180 7.5.4 Other functions of NBEAL2 in megakaryocytes and platelets ...... 180

References ...... 182

List of Tables

Table 4.1. List of potential NBEAL2 N-terminal interacting partners from yeast-two-hybrid screens ...... 94 Table 4.2. List of potential NBEAL2 C-terminal interacting partners from yeast-two-hybrid screens ...... 95 Table 4.3. Potential NBEAL2 binding partners obtained from 3XFlag-NBEAL2 IP mass spectrometry with DAMI cells ...... 97

List of Figures

Figure 1.1. The roles of platelets in hemostasis ...... 5 Figure 1.2. Thrombosis impedes blood flow and alters hemodynamics ...... 9 Figure 1.3. Leukocyte-platelet interaction contributes to the development of the atherosclerotic plaque ...... 12 Figure 1.4. The presence of atherosclerotic plaque increases the incidences of severe thrombosis ...... 13 Figure 1.5. Megakaryocyte development and maturation ...... 17 Figure 1.6. Visualization of a proplatelet structure ...... 18 Figure 1.7. Anatomy of a platelet ...... 19 Figure 1.8. Ultrastructure of platelets from ARC and GPS patients ...... 33 Figure 1.9. Ultrastructure of platelets from ARC and GPS patients ...... 34 Figure 1.10. Model of α-granule biogenesis in megakaryocytes ...... 37 Figure 1.11. Diagram of NBEAL2 displaying homology domains and GPS-causing mutations ...... 41 Figure 1.12. Schematic of the domains of BDCP ...... 41 Figure 3.1. NBEAL2 is capable of oligomerization ...... 65 Figure 3.2. Validation of NBEAL2 antibody and Nbeal2-/- mouse ...... 66 Figure 3.3. Schematic of NBEAL2 constructs used to generate stable cell lines ...... 67 Figure 3.4. Generation of stable DAMI cell lines that are expressing NBEAL2 ...... 68 Figure 3.5. Validation of NBEAL2 antibody for immunofluorescence ...... 69 Figure 3.6. Validation of anti-NBEAL2 antibody for immunostaining...... 70 Figure 3.7. Schematic for extracting cytosolic and membrane fractions from cells ...... 71 Figure 3.8. NBEAL2 is primarily cytosolic in DAMI cells ...... 72 Figure 3.9. Fractionation of primary human megakaryocytes ...... 74 Figure 3.10. NBEAL2 colocalizes with P-selectin in primary human megakaryocytes ...... 76 Figure 3.11. NBEAL2 interacts with P-selectin ...... 78 Figure 3.12. NBEAL2 interacts with P-selectin in cultured primary human megakaryocytes . 79 Figure 3.13. Schematic of the NBEAL2 PH-BEACH construct ...... 80 Figure 3.14. Expression and purification of PH-BEACH ...... 81 Figure 3.15. Purification of PH-BEACH with cation exchange chromatography ...... 82 Figure 3.16. Purification of PH-BEACH with gel-filtration chromatography ...... 83 Figure 3.17. Strategies to improve PH-BEACH protein quality ...... 84

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Figure 3.18. Expression of PH-BEACH in mammalian cells ...... 86 Figure 4.1. Schematic of NBEAL2 constructs used for the yeast-two-hybrid screens ...... 93 Figure 4.2. Expression of NBEAL2 bait constructs ...... 93 Figure 4.3. Confirmation of yeast-two-hybrid hits ...... 96 Figure 4.4. NBEAL2 interacts with FLNA ...... 98 Figure 4.5. NBEAL2 interacts with 14-3-3 proteins at the endogenous level ...... 100 Figure 4.6. NBEAL2 interacts with SEC22B ...... 102 Figure 4.7. Domain diagrams of NBEAL2 and SEC22B ...... 103 Figure 4.8. Confirmation of NBEAL2-SEC22B interaction ...... 104 Figure 4.9. Mapping the binding regions of NBEAL2 to SEC22B ...... 105 Figure 4.10. NBEAL2 contains inhibitory elements that prevent its interaction with SEC22B ...... 107 Figure 4.11. Defining the binding region of NBEAL2 to SEC22B ...... 108 Figure 4.12. Expression of NBEAL2 is dependent on SEC22B ...... 109 Figure 4.13. SEC22B is required for α-granule formation in imMKCL cells ...... 111 Figure 4.14. NBEAL2 E1833K and R1839C abrogates NBEAL2-SEC22B interaction ...... 113 Figure 5.1. Blood film and ultrastructure abnormalities of Nbeal2-/- mouse platelets ...... 123 Figure 5.2. Granule cargo proteins, P-selectin and NBEAL2 in platelets and megakaryocytes ...... 125 Figure 5.3. P-selectin in resting and thrombin-activated Nbeal2-/- platelets ...... 126 Figure 5.4. Impaired Nbeal2-/- mouse platelet function in vitro and in vivo ...... 128 Figure 5.5. Abnormal ultrastructure of Nbeal2-/- bone marrow megakaryocytes ...... 129 Figure 5.6. Nbeal2-/- megakaryocytes showed reduced survival or differentiation ...... 131 Figure 5.7. Abnormal Nbeal2-/- megakaryocyte development in populations of cultured bone marrow cells ...... 133 Figure 5.8. Delayed maturation of native Nbeal2-/- megakaryocytes ...... 134 Figure 5.9. Abnormal VWF distribution in Nbeal2-/- megakaryocytes ...... 135 Figure 5.10. Von Willebrand factor is not trafficked to proplatelets in cultured Nbeal2-/- megakaryocytes ...... 136 Figure 5.11. Cultured Nbeal2-/- megakaryocyte in spread phase ...... 137 Figure 5.12. Cultured Nbeal2-/- megakaryocyte with proplatelets ...... 138 Figure 5.13. Reduced presence of TSP1 in Nbeal2-/- megakaryocytes ...... 139 Figure 5.14. TSP1 is not trafficked to proplatelets in cultured Nbeal2-/- megakaryocytes .... 140 Figure 5.15. TSP1 secretion by Nbeal2-/- megakaryocytes ...... 141

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Figure 6.1. Abnormal VWF trafficking in Nbeal2-/- megakaryocytes ...... 148 Figure 6.2. FGN level is reduced in Nbeal2-/- platelets ...... 149 Figure 6.3. Abnormal VWF and FGN localization in Nbeal2-/- megakaryocytes ...... 150 Figure 6.4. Transgenic expression of GFP-NBEAL2 in Nbeal2-/- mice ...... 151 Figure 6.5. The expression of the NBEAL2 transgene in Nbeal2-/- mice did not propagate . 152 Figure 6.6. Validating the expression of NBEAL2 in Nbeal2-/- mice ...... 153 Figure 6.7. Differences in fibrinogen endocytosis by WT and Nbeal2-/- megakaryocytes ... 154 Figure 6.8. Endocytosed fibrinogen is associated with multivesicular body-like structures in WT megakaryocytes ...... 154 Figure 6.9. Differences in fibrinogen endocytosis by WT and Nbeal2-/- megakaryocytes .... 156 Figure 6.10. Endocytosed fibrinogen is not aberrantly degraded in Nbeal2-/- megakaryocytes ...... 157 Figure 6.11. Accelerated release of endocytosed fibrinogen by Nbeal2-/- megakaryocytes . 158 Figure 6.12. Endocytosed fibrinogen is abnormally distributed in Nbeal2-/- megakaryocytes ...... 161 Figure 6.13. Differential trafficking of endocytosed fibrinogen by WT and Nbeal2-/- megakaryocytes ...... 162 Figure 6.14. Convergence of endocytosed fibrinogen in WT megakaryocytes ...... 163 Figure 6.15. Abnormal trafficking of VWF by Nbeal2-/- megakaryocytes ...... 165 Figure 6.16. Abnormal trafficking of endogenously-synthesized and endocytosed granule cargo by Nbeal2-/- megakaryocytes ...... 166 Figure 6.17. α-granule externalization is not the cause of VWF release in Nbeal2-/- megakaryocytes ...... 167 Figure 7.1. Model of the role of NBEAL2 in α-granule cargo trafficking and maturation ...... 176

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List of Abbreviations

AcLDL Acetylated LDL ADP Adenosine-5'-diphosphate ARC Arthrogryposis, Renal Dysfunction and Cholestasis AP Affinity purification ARF6 ADP-ribosylation factor 6 ARM Armadillo ATP Adenosine-5'-triphosphate BEACH Beige and CHS Bchs Blue cheese BDCP BEACH domain containing proteins BIT Bovine serum albumin, insulin, and transferrin BLOC Biogenesis of lysosome-related organelles complex BSA Bovine serum albumin CD Cluster of differentiation cDNA Complementary DNA CHS Chediak-Higashi syndrome ConA Conacavalin A-like lectin CORVET Class C core vacuole/endosome tethering CRISPR Clustered regularly interspaced short palindromic repeats DMEM Dulbecco's Modified Eagle Medium DMS Demaracation membrane system DNA Deoxyribonucleic acid DTS Dense tubular system ECM Extracellular matrices ER Endoplasmic reticulum ERGIC Endoplasmic reticulum-Golgi intermediate compartments ENA-78 Epithelial cell-derived neutrophil-activating 78 FGF Fibroblast growth factor FOG1 Friend of GATA1 FGN Fibrinogen GAPDH Glyceraldehyde 3-phosphate dehydrogenase GEF Guanine nucleotide exchange factor GFP Green fluorescent protein GP Glycoprotein GPS Gray platelet syndrome GS Griscelli syndrome GTP Guanosine-5'-triphosphate HMWK High-molecular-weight kininogen HOPS Homotypic fusion and vacuole protein sorting HPS Hermansky-Pudlak syndrome HRP Horseradish peroxidase HS Horse serum HSC Hematopoietic stem cells IB Immunoblot IF Immunofluorescence IP Immunoprecipitation IMDM Iscove’s Modified Dulbecco’s Medium ImMKCL Immortalized megakaryocyte progenitor cell IMS Invaginated membrane system

LAMP Lysosomal-associated membrane protein LDL Low density lipoproteins LRBA Lipopolysaccharide-responsive, beige-like anchor protein LRO Lysosome-related organelle LYST Lysosomal trafficking regulator KO Knockout MCS Membrane contact sites MK Megakaryocyte mRNA Messenger RNA MMP Matrix metalloproteinases MS Mass spectrometry MVB Multivesicular body NBEA Neurobeachin NBEAL Neurobeachin-like NSMAF Neutral sphingomyelinase activation-associated factor OCS Open canalicular system OxLDL Oxidized LDL PAI-1 Plasminogen activator inhibitor-1 PBS Phosphate buffered saline PCR Polymerase chain reaction PDGF Platelet-derived growth factor PF4 Platelet factor 4 PFA Paraformaldehyde PH Pleckstrin homology PK Prekallikrein PM Plasma membrane PRP Platelet-rich plasma PS Phosphatidyl serine PSGL-1 P-selectin glycoprotein ligand-1 MCP-3 Monocyte chemoattractant protein-3 NAP-2 Neutrophil-activating peptide 2 SEC-MALS Size exclusion chromatography with inline multi-angle light scattering SDF1 Stromal cell-derived factor 1 SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis SNARE Soluble N-ethylmaleimide-sensitive factor attachment protein receptor TAFI Thrombin activatable fibrinolysis inhibitor TEM Transmission electron microscopy TF Tissue factor TGF-β Transforming growth factor beta TLR Toll-like receptors TM Transmembrane tPA Tissue plasminogen activator TPO Thrombopoietin TSP1 Thrombospondin-1 VAMP Vesicle-associated membrane protein VAPA Vesicle-associated membrane protein-associated protein A VSMC Vascular smooth muscle cells VEGF Vascular endothelial growth factor VWF von Willebrand factor WDFY WD and FYVE zinc finger domain containing protein WDR81 WD repeat domain 81

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WT Wild-type

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Chapter 1

Introduction

1.1 Platelet functions 1.1.1 Overview

Platelets are small (2-3 μm diameter) discoid anucleate blood cells derived from megakaryocytes (MKs), which in adult mammals reside primarily in the bone marrow. Platelets are the cellular effectors of hemostasis (Hartwig and Italiano 2003), which is essential for the maintenance of vasculature integrity. Platelets adhere to sites of vascular damage, where they become activated and begin to aggregate to form a seal known as the hemostatic or platelet plug, which serves as an initial seal for the wound that prevents further blood loss. Central to this process is the release of the contents of secretory granules, α- and δ-granules in particular, during platelet activation. Secreted molecules activate other platelets in the vicinity, recruiting them to the injury site, a positive feedback that is critical for the establishment of the hemostatic plug (Patel, Hartwig et al. 2005). Activated platelets also provide procoagulant surfaces for the generation of thrombin that catalyzes the production of fibrin, which combines with aggregated platelets to produce a strong and stable hemostatic clot (Sidelmann, Gram et al. 2000). Impairments in the formation or function of platelet secretory granules are associated with bleeding disorders. For example, α-granule deficiencies are linked with Arthrogryposis, Renal tubular dysfunction, and Cholestasis (ARC) syndrome and Gray Platelet Syndrome (GPS), while δ-granule deficiencies are associated with bleeding disorders including Hermansky-Pudlak syndrome, Chediak-Higashi syndrome and Griscelli syndrome (Diz-Kucukkaya 2013).

The role of platelets in hemostasis is well characterized, and recent advances have shed light on other functions of platelets, many associated with the wide assortment of bioactive molecules stored within their secretory granules that can mediate physiological processes such as host immunity and wound healing. The vast array of clotting proteins, growth factors, chemokines and angiogenic factors carried by platelets have also been implicated in the pathogenesis of thrombosis, atherosclerosis, angiogenesis and malignancies (Nurden, Nurden et al. 2008, Smyth, McEver et al. 2009, Ali, Wuescher et al. 2015). The implication of platelets in a diverse array of processes owing to the multitude of bioactive molecules stored within their secretory granules makes it important to understand the biogenesis of those granules, yet many aspects remain undefined. Elucidating the mechanisms of α- and δ- granule development, function and regulation will also advance understanding of platelet disorders.

1.1.2 Roles of platelets in hemostasis

Platelets are key to the initial response to sealing vessel wounds, or primary hemostasis. When vascular injury occurs endothelial cells are ruptured or removed to expose the subendothelial matrix, which is rich in collagen, a potent platelet activator (Chen and Lopez 2005). Initial adhesion of platelets to the subendotheium occurs via the interaction between platelet integrin GPIa-IIa (integrin α2β1) and GPVI with collagen (Nieswandt, Brakebusch et al. 2001, Sarratt, Chen et al. 2005). These interactions are sufficient for platelets to adhere to the subendothelial matrix under low shear conditions. Under high shear rates, additional interactions between platelet glycoprotein receptor complex GPIb-IX-V, von Willebrand factor (VWF) and collagen are required to stabilize adhesion. In this process, plasma VWF first recognizes and binds to the exposed collagen, thereby becoming immobilized onto the subendothium. The high shear rates of the circulation causes the immobilized VWF to undergo a conformational change, revealing the A1 domain that binds to GPIbα subunit of the platelet integrin complex, GPIb-IX-V. Platelets move along the subendothelium as a result of the interactions between GPIb-IX-V, VWF and collagen, halting its speed and eventually bringing platelets to a firm adherence to the subendotheium (Cranmer, Ulsemer et al. 1999, Arthur, Gardiner et al. 2005, Andrews and Berndt 2008). Once this occurs, the binding of platelet integrin complex α2β1 to collagen further stabilizes the adhesion and the binding of platelet glycoprotein receptor GPVI to collagen activates the platelets via intracellular signal transduction (Nieswandt, Brakebusch et al. 2001, Sarratt, Chen et al. 2005).

Activated platelets change shape as they become larger and produce multiple cytoplasmic extensions that allow them to spread over the subendothelium. This creates a larger surface area of contact for further adhesion and aggregation (Maxwell, Westein et al. 2007). Platelet aggregation is achieved via GPIIb-IIIa (integrin αIIbβ3) linkages, and this process is essential to the formation and growth of the hemostatic plug. Integrin αIIbβ3 is normally inactive on resting platelets, but following activation αIIbβ3 changes conformation and becomes capable of binding to VWF and fibrinogen (FGN). Activated αIIbβ3 receptors bind to VWF immobilized on subendothelial collagen, promoting a firm adhesion of the activated platelets to the injury site. FGN bridges activated platelets by binding to activated αIIbβ3 receptors, linking platelets together and strengthening platelet aggregates. (Fullard 2004, Peyvandi, Garagiola et al. 2011, Schneider 2011).

Simultaneously, activated platelets undergo degranulation and release the contents of secretory granules to the surrounding environment. These molecules include platelet agonists such as prothrombin (converted into active thrombin via secondary hemostasis) and adenosine diphosphate (ADP), which activates additional platelets and recruits them to the injury site. In addition to their presence in the plasma, FGN and VWF are also stored within α- granules. α-granule secretion generates a local concentration of clotting proteins, including FGN and VWF, at the site of injury, which aids in the formation and stabilization of the hemostatic plug (Blair and Flaumenhaft 2009, Golebiewska and Poole 2015). As platelets continue to aggregate, the hemostatic plug grows in size and eventually seals the blood vessel break.

While primary hemostasis is efficient in creating an initial seal to the vessel break, further stabilization is needed to strengthen the blockage as the hemostatic plug grows. This process is facilitated by secondary hemostasis, also known as coagulation. Secondary hemostasis occurs simultaneously with primary hemostasis and the end result is the incorporation of fibrins into the primary hemostatic plug, producing a fibrin clot that is highly stabilized and is resistant to physical shears and chemical alterations. This strengthens the integrity of the primary plug and aids in the binding of the plug to the injury site (Gale 2011).

Coagulation can be initiated via either the intrinsic or extrinsic pathway. The intrinsic pathway relies on the contact between circulatory Factor XII (FXII), plasma high-molecular-weight kininogen (HMWK) and prekallikrein (PK) on subendothelial collagen. This converts inactive FXII to the active FXIIa, leading to a cascade of events that eventually activates Factor X (FX) into FXa (Mutch 2011, Renne, Schmaier et al. 2012). Extrinsic pathway is initiated by the binding of Factor VII (FVII) in circulation to the exposed subendothelial tissue factor (TF) following vasculature damage, forming an activated complex of TF-FVIIa. TF-FVIIa then activates FX into FXa (Camerer, Huang et al. 2000, Norledge, Petrovan et al. 2003). FXa, the end product from the activation of both the intrinsic and extrinsic pathway, associates with Factor Va (FVa) to form the prothrombinase complex, which ultimately converts the inactive plasma prothrombin into active thrombin (Schuijt, Bakhtiari et al. 2013, Podoplelova, Sveshnikova et al. 2016). The assembly and activity of the prothrombinase complex requires a negatively charged phospholipid membrane surface (Billy, Willems et al. 1995). One such surface is provided by the activated platelet where, during platelet activation, phosphatidyl serine (PS) which is rich on the inner membrane leaflet is flipped to the exterior, creating a

4 procoagulant surface (Lentz 2003). Following activation, thrombin cleaves soluble plasma FGN into insoluble fibrin on the surfaces of activated platelets. Crosslinking of these fibrins by activated factor XIII creates a mesh that forms the basis of a stable fibrin clot (Standeven, Carter et al. 2007, Wolberg and Campbell 2008). Activated platelets also secrete FGN and prothrombin that are housed in their α-granules to amplify the local concentration of these proteins at the site of injury (Golebiewska and Poole 2015).

The coagulation cascade is highly regulated in order to maintain coordinated and localized clotting, as well as to prevent undesirable clotting outside of the site of tissue damage. For example, plasma tissue factor pathway inhibitor (TFPI) inactivates TF-FVIIa and FXa, and plasma antithrombin inhibits thrombin, FXa and other components of the coagulation cascade. These regulations limit the activity of the coagulation factors at the site of vessel damage (Mackman 2009). As the wound heals, the fibrin clot is resorbed via plasmin-mediated fibrinolysis (Laurens, Koolwijk et al. 2006). A schematic of the processes of hemostasis is shown in Fig. 1.1.

Figure 1.1. The roles of platelets in hemostasis. Vascular injury induces a break in the vessel wall and exposes the subendothelial matrix. (A) The binding of circulating platelets to subendothelial collagen initiates primary hemostasis. The interaction between collagen, VWF and platelet integrin complex, GPIb-IX-V, promotes the firm adhesion of platelets to the subendothelial matrix and the binding of collagen to platelet integrin, GPVI, activates the platelets. Activated platelets display morphology changes and undergo degranulation. This results in a cascade response that recruits additional platelets to the injury site, induces aggregation and forms the primary hemostatic plug. (B) The primary hemostatic plug is strengthened by the incorporation of fibrin, the product of secondary hemostasis. The exposure of triggers such as TF in the subendothelial matrix activates the coagulation cascade. Aggregating platelets provides a phospholipid surface that is required for the assembly and activation of the coagulation complexes. The coagulation cascade converts prothrombin into thrombin, which in turn cleaves FGN into fibrin. Polymerization and crosslinking of fibrin strands stabilizes the hemostatic plug and produces a fibrin clot. Figure adapted and reprinted with permission (Cito, Mazzeo et al. 2013).

1.1.3 Other physiological roles of platelets

Platelets are traditionally known for their roles in hemostasis and thrombosis. The former, a physiological function to prevent bleeding and the latter, a pathological condition that impedes and obstructs blood flow as clots are being deposited within the blood vessel. Recent advances have uncovered and characterized novel physiological roles for platelets. Most notably, inflammation, host immunity and wound healing. These functions of platelets are largely attributed to the molecules stored within their secretory granules. For example, platelet derived growth factor (PDGF), fibroblast growth factor (FGF) and transforming growth factor beta (TGF-β) are platelet α-granule proteins that are known to promote wound healing. Upon release from platelets, these factors stimulate the growth and proliferation of endothelial cells and fibroblasts and promote their migration into the injury site (Laurens, Koolwijk et al. 2006).

Inflammation is a physiological response that removes irritants or pathogens and repairs tissues following tissue damage. During inflammation, damaged tissues attract immune cells via chemokine release. Circulatory immune cells can either adhere to the vascular injury site to become activated, or tether to activated endothelium to gain access to the site of tissue damage via transmigration. Platelets produce and store pro-inflammatory factors within their secretory granules. In the event of vascular or tissue injury, platelets become activated at or near the site of injury and release inflammatory factors through degranulation. These cytokines and chemokines then promote a local inflammatory response cascade by recruiting, activating and influencing the functions of immune cells at the site of damage (Stokes and Granger 2012). For example, α-granule constituent interleukin-8 and monocyte chemoattractant protein-3 (MCP-3) recruit leukocytes and monocytes to the site of injury; platelet factor 4 (PF4), neutrophil-activating peptide 2 (NAP-2) and epithelial cell-derived neutrophil-activating 78 (ENA-78) activate leukocytes and promote their adhesion to the injury site; stromal cell-derived factor 1 (SDF1) regulates monocyte activation and differentiation (Weyrich and Zimmerman 2004, Semple, Italiano et al. 2011, Chatterjee, von Ungern- Sternberg et al. 2015). In addition to cytokine and chemokine release, platelets can physically interact with leukocytes to facilitate inflammatory responses. During platelet degranulation, α- granules are translocated to the plasma membrane for exocytosis via vesicular fusion. As such, while α-granule cargo proteins are secreted, α-granule membrane constituents become incorporated within the platelet plasma membrane, resulting in the surface expression of granule membrane proteins such as P-selectin (Fitch-Tewfik and Flaumenhaft 2013). P-

6 selectin subsequently binds to P-selectin glycoprotein ligand-1 (PSGL-1) on leukocytes. Binding of P-selectin to PSGL-1 activates leukocytes. This enhances the adhesion of leukocytes to the injury site and elevates the release of chemokines from leukocytes, leading to an amplified inflammatory response that promotes the recruitment of additional immune cells to the injury site (Smyth, McEver et al. 2009, Vieira-de-Abreu, Campbell et al. 2012, Ali, Wuescher et al. 2015). Other platelet membrane receptors are also known to activate or enhance the functions of immune cells. For example, the binding of platelet surface protein, CD40 ligand (CD40L, also known as CD154) to neutrophil’s CD40 receptor activates the neutrophils and promotes their adhesion to the inflammation site (Henn, Steinbach et al. 2001).

In addition to platelets influencing the functions of leukocytes, activated platelets can adhere to inflamed or activated vascular endothelial cells to further promote inflammation. Activated endothelial cells express adhesive proteins on the cell surface which facilitates the tethering and adhesion of platelets. For example, platelet glycoprotein receptor complex, GPIb-IX-V, binds to P-selectin on the surface of activated endothelial cells (Romo, Dong et al. 1999); activated platelet integrin αIIbβ3, binds to endothelial αvβ3 integrin via FGN linkage (Li, Podolsky et al. 1996, Bombeli, Schwartz et al. 1998); the binding of platelet CD40L to endothelial CD40 induces endothelial activation (Henn, Slupsky et al. 1998). The interaction between platelets and endothelial cells activates endothelial cells and enhances their expression and release of pro-inflammatory cytokines and chemokines. Similarly, activated platelets also release pro-inflammatory molecules through degranulation. The release of pro- inflammatory molecules from both the inflamed endothelial cells and activated platelets recruit leukocytes to the site, facilitate leukocyte transmigration across the endothelium and propagate the inflammation process (Kaplan and Jackson 2011).

Platelets are intimately involved with host immunity and prevention of infection. For example, platelets can activate dendritic cells (DC) through CD154-CD40 interaction to promote adaptive immune responses (Henn, Steinbach et al. 2001). In addition to influencing the functions of immune cells, platelets are capable of recognizing and destroying infectious agents such as bacteria. Platelets express toll-like receptors (TLRs), a class of recognition receptors also expressed by leukocytes and macrophages, enabling them to identify bacteria (Andonegui, Kerfoot et al. 2005). Upon recognition by TLRs, platelets undergo degranulation and destroy the bacteria by releasing microbicidal proteins such as thrombocidins.

Alternatively, platelets can destroy bacteria by internalization or by first trapping them through aggregation, then recruiting phagocytes such as macrophages to eliminate the bacteria (McNicol and Israels 2008, Smyth, McEver et al. 2009). Platelets are also known to participate in the activation of the complement pathway, leading to increased levels of soluble C3a and C5a to promote the chemotaxis and activation of immune cells. Platelets can activate or amplify the intensity of the complement cascade in different manners: activated platelets provide a PS surface that is required for the assembly of C5b, C6 and C7; P-selectin on the surface of activated platelets can bind to C3b, leading to the activation of C3 convertase, which in turn promotes the production of C3a from C3 (Del Conde, Cruz et al. 2005, Hou, Carrim et al. 2015, Afshar-Kharghan 2017). As more functions of platelets are being discovered, it is clear that platelets contribute to a wide variety of biological processes and their roles are not limited to hemostasis and thrombosis.

1.1.4 Pathological roles of platelets 1.1.4.1 Platelets, atheroscelerosis and thrombosis

Platelets are crucial components of hemostasis and as previously described, also participate in a diverse variety of biological processes. Due to this diverse role, dysregulated platelet function can lead to pathological thrombotic conditions. Platelets have traditionally been implicated in acute coronary syndrome and stroke due to vascular occlusion of arteries to the heart and brain respectively (Furie and Furie 2008).Thrombosis is a pathological condition where the formation, growth or resorption of the clot becomes misregulated and the physical presence of the clot obstructs the flow of blood within the blood vessel (Packham 1994). While the severity of thrombosis may vary, thrombi that dislodge from the vessel wall and travel towards the pulmonary artery can cause vessel occlusion in the lung. The blockage of blood vessels in the lung leads to pulmonary embolism, a life-threatening condition that manifests in lung dysfunction, which results in the deprivation of oxygen in all body tissues (Chung, Connor et al. 2007).

Typically, fibrin clot or thrombus is broken down by plasmin through fibrinolysis. Fibrinolysis cuts fibrin via proteolytic cleavage, leading to the destabilization and disintegration of the thrombus (Chapin and Hajjar 2015). Fibrinolysis is however countered by the activity of platelets during thrombus formation. To promote the stabilization of the thrombus and to prevent premature thrombus removal, platelets secrete anti-fibrinolytic agents such as α- granule proteins plasminogen activator inhibitor-1 (PAI-1) and thrombin activatable fibrinolysis

8 inhibitor (TAFI) upon activation (Mosnier, Buijtenhuijs et al. 2003, Mutch, Thomas et al. 2007). As wound healing progresses, the thrombus is removed as proliferating fibroblasts and endothelial cells migrate into the site of injury and secrete pro-fibrinolysis factors such as tissue plasminogen activator (tPA) and urokinase, both which activates plasminogen into plasmin (Schafer, Maier et al. 1994, Chan, Duszczyszyn et al. 2001). A balance is struck between pro- and anti-fibrinolysis factors to determine the appropriate time for thrombus removal. It is clear that not only do platelets play key roles in the formation of the thrombus during hemostasis, platelets also participate in regulating thrombus clearance. Aberrant and hyperactive platelet activities are characterized in severe cases of thrombosis (Cay, Ipek et al. 2012, Zhang, Ye et al. 2012). An illustration of a thrombotic event is shown in Figure 1.2.

Figure 1.2. Thrombosis impedes blood flow and alters hemodynamics. Thrombosis occurs when the formation of a blood clot obstructs the blood flow. While the fibrin polymerization stabilizes the primary hemostatic plug, it can also trap blood cells within the thrombus. The fibrin clot is dissociated as proliferating fibroblast and endothelial cells migrate into the site. Fibroblasts and endothelial cells secrete and express an assortment of proteins to modulate this process, including the secretion of tissue plasminogen activator (tPA) for fibrinolysis and the expression of surface receptor thrombomodulin for the inhibition of the coagulation cascade. When a thrombus grows excessively or when fibrinolysis is impaired, the thrombus can impede the blood flow within the vessel and change the hemodynamics. Severe thrombosis can lead to vessel occlusion and causes ischemic episodes. Further, an unresolved thrombus may break off from its site of formation and through migration, causes the occlusion of another blood vessel. This is known as thromboembolism. Figure adapted and reprinted with permission (Cito, Mazzeo et al. 2013).

Platelet function is intimately associated with coronary heart disease and stroke, characterized by atherothrombosis, a condition involving the complex interplays between inflammation and thrombotic events. Atherosclerosis is a pathological condition resulting in narrowing of arteries due to plaque accumulation in the subendothelial space. As the plaque grows, blood supplies to body tissues decrease, leading to ischemic stroke and myocardial infarction when arteries supplying blood to the brain and heart are completely obstructed. Atherosclerotic plaques are mainly composed of vascular smooth muscle cells (VSMCs), macrophages and foam cells (Bennett, Sinha et al. 2016). The initiation and growth of the

9 plaques stem from tissue damage which activates vascular endothelial cells, both of which generate inflammation triggers (Wallach, Kang et al. 2014). In addition to tissue damage, the presence of reactive oxygen species (ROS) and modified lipoproteins such as acetylated low density lipoprotein (AcLDL) and oxidized low density lipoproteins (OxLDL) are also potent triggers of endothelial inflammation (Kaplan and Jackson 2011, Gradinaru, Borsa et al. 2015).

Activation of endothelial cells enhances the permeability of the endothelium to leukocyte infiltration and promotes the expression and release of pro-inflammatory cytokines and chemokines (Gros, Ollivier et al. 2014). The generation of inflammatory signals trigger and enable circulating monocytes to adhere and transmigrate across the arterial endothelium into the subendothelial space. Within the subendothelial space, inflammation signals further promote monocytes to differentiate into macrophages and upon ingestion and metabolism of modified lipoproteins, including AcLDL and OxLDL, transform into lipid-rich foam cells (Hayden, Brachova et al. 2002, Shi and Pamer 2011). Foam cells subsequently release pro- inflammatory cytokines and chemokines in addition to reactive oxygen species (ROS) and growth factors. These molecules propagate the inflammatory signals, recruit additional immune cells to the site and stimulate the proliferation of VSMCs (Yu, Fu et al. 2013). VSMCs similarly transform into foam cells upon ingestion of modified lipoproteins and further propagate the inflammation process (Chaabane, Coen et al. 2014). As the numbers of macrophages, foam cells and dead cells increase, the plaque grows, hardens and pushes against the arterial lumen. The fibrous cap, a layer of connective tissues that separates the atherosclerotic lesion from the arterial lumen, becomes unstable as the plaque expands and eventually ruptures or erodes. Disruption of the arterial vessel walls recruit platelets to the site and consequentially, thrombosis occurs (Kaplan and Jackson 2011, Badimon and Vilahur 2014). The thrombus that forms within the arterial lumen occludes the artery, effectively reducing or preventing the supply of blood to organs and tissues. Thromboembolism can also occur if the thrombus dislodges from the vessel wall and migrates to block additional blood vessels.

Platelets are potent mediators of atherothrombosis. Reports have shown that platelet activities are enhanced or altered in cases of atherothrombosis and are more likely to interact with leukocytes, thereby promoting the activation and migration of leukocytes into the atherosclerotic sites (Totani and Evangelista 2010). Platelet’s role in atherothrombosis are linked to their functions in hemostasis and inflammation. As previously mentioned, upon

10 activation by tissue damage, platelets can facilitate the propagation of inflammation by the release of pro-inflammatory factors through degranulation and by enhancing the functions of activated vascular endothelial cells (Li, Podolsky et al. 1996, Gawaz, Neumann et al. 1997, Bombeli, Schwartz et al. 1998, Romo, Dong et al. 1999). The binding of platelets to activated endothelial cells increases the permeability of the endothelium to leukocyte infiltration and promotes the release of pro-inflammatory molecules from the endothelial cells. The pro- inflammatory factors released by both platelets and endothelial cells recruit and activate leukocytes for adhesion and transmigration into the subendothelial space, and promotes the formation of foam cells (Davi and Patrono 2007, Kapoor 2008, Langer and Gawaz 2008, May, Seizer et al. 2008, Quinn, Henriques et al. 2011, Gros, Ollivier et al. 2014). During this process, the activities of leukocytes are further enhanced by platelet-leukocyte interactions via surface receptor interactions. A key interaction is the ligation of platelet surface P-selectin with leukocyte PSGL-1. This activates the leukocytes and enhances their ability to adhere and transmigrate across the endothelium (McEver and Cummings 1997, Frenette, Denis et al. 2000). An illustration of platelet-leukocyte interaction in atherogenesis is shown in Fig. 1.3.

The cytokines and chemokines released from activated platelets and endothelial cells also activate other resting platelets in the circulation, thereby propelling the cycles of inflammation and amplifying the inflammatory response (Wagner and Frenette 2008). Thrombosis that occurs following the rupture of the atherosclerotic lesion is also a hemostasis process that is dependent on the activity of platelets. During the formation of the thrombus, leukocytes are also recruited to the site. This process is driven by platelet degranulation and further aggravating conditions (Saha, Humphries et al. 2011, Swystun and Liaw 2016). The presence of an underlying atherosclerotic plaque in the blood vessel also increases the risk of developing severe cases of thrombosis as the thrombus that is formed following plaque rupture can only grow inside the arterial lumen (Fig. 1.4). Clearly, platelets play key roles in promoting and propagating inflammation response and facilitating atherothrombosis.

Normally, blood vessels are protected from atherosclerotic events as the adhesion of monocytes to the endothelium is inhibited and regulated by vascular endothelial cells. Specifically, endothelial cells produce ecto-ADPase, nitric oxide and prostacyclin, all of which are potent in inhibiting the activation and adhesion of platelets and leukocytes (Marcus, Broekman et al. 1997, Jin, Voetsch et al. 2005, Kaplan and Jackson 2011). However, in the event of dysfunctional endothelium or chronic inflammation, these functions are lost or

11 impaired, either due to defects or as a natural physiological response to inflammation. Alteration of platelet functions by exogenous factors such as OxLDL can also negate the endothelium’s protective mechanisms (Gradinaru, Borsa et al. 2015). Platelets that are activated via OxLDL internalization are shown to be capable of adhering and activating endothelial cells directly despite the absence of pre-existing inflammation. In addition, while it has been shown that through P-selectin and PSGL-1 that interaction that platelets can promote the differentiation of CD34+ HSCs into endothelial cells and foam cells, this relationship is worsened by the presence of OxLDL. When compared with normal platelets, incubation of CD34+ HSCs with OxLDL-laden platelets drastically increases foam cell differentiation while limiting the endothelial lineage (Daub, Langer et al. 2006, Daub, Seizer et al. 2010). Altogether, it is clear that the complex interplays between vascular endothelial cells, leukocytes and platelets are critical to the initiation and progression of atherothrombosis.

Figure 1.3. Leukocyte-platelet interaction contributes to the development of the atherosclerotic plaque. The transmigration of leukocyte across the endothelium is a physiological response to inflammation and is also a prerequisite for the development of the atherosclerotic plaque. Leukocytes are first recruited to the endothelium via chemokine and cytokines released by activated platelets and endothelial cells. Ligation of platelet P-selectin to leukocyte PSGL-1 activates the leukocytes and promotes adhesion. After the initial tethering, additional receptor interactions stabilize the adhesion and transmigration of the leukocytes. Monocytes that transmigrate across the endothelium internalizes OxLDL and transforms into foam cells, contributing to the formation of the atherosclerotic plaque. During this entire process, platelets continuously secrete molecules that enhances monocyte adhesion and differentiation (e.g. RANTES). Figure adapted and reprinted with permission (Gawaz, Langer et al. 2005).

Figure 1.4. The presence of atherosclerotic plaque increases the incidences of severe thrombosis. Under normal conditions, the formation of fibrin clots does not usually impede blood flow as the thrombus tends to grow into the extraluminal space instead of towards the arterial lumen. The presence of the atherosclerotic plaque, however, prevents the growth of the fibrin clot in the extraluminal direction, and thus encourages the formation of an intraluminal thrombus. Severe thrombosis can cause the occlusion of the artery and results in ischemic episodes. The intraluminal thrombus is also then exposed to the physical and chemical shears of the blood circulation and thereby increasing the risks of thromboembolism. Figure adapted and reprinted with permission (Jackson 2011).

1.4.4.2 Platelets and cancer

One of the first concrete lines of evidence that implicated platelets in cancer was reported in 1968 when it was revealed that neuraminidase-induced thrombocytopenia in mice protects them against tumor growth and metastasis (Gasic, Gasic et al. 1968). Since then, reports have identified increased platelet activity in cancer patients, further suggesting the likelihood that platelets may play active roles in cancer development. The hallmarks of cancer cells are their abilities to metastasize, circumvent apoptosis and induced cell death, maintain boundless proliferation and growth and promote angiogenesis. Platelet α-granules contain a vast array of proteins that are thought to be capable of promoting tumorigenesis. Supporting

13 this notion, it was shown that mice that lack platelet α-granules are protected from tumor growth and metastasis, discerning the importance of α-granules in tumorigenesis (Guerrero, Bennett et al. 2014). This is not unexpected as platelet α-granules are rich in growth factors that may contribute to the sustainability of tumor cells. This is exemplified by the increased carcinoma cell proliferation when the cells are incubated with platelet releasate (Janowska- Wieczorek, Wysoczynski et al. 2005).

Cancer metastasis is characterized by the growth of secondary tumors that originated from a distant primary tumor. Tumor cells typically migrate from their primary to their secondary sites via blood or lymph vessel transport and during this process, tumor cells must first invade the extracellular matrices (ECM) that encloses the primary site and evade immune surveillances once they are in the circulation. Carcinoma cells that are incubated with platelet releasates expressed higher levels of matrix metalloproteinases (MMPs) and showed increased invasive properties (Janowska-Wieczorek, Wysoczynski et al. 2005). In blood vessels, metastasizing cancer cells can recruit platelets to form a barrier that encloses the tumor cell within, protecting the cell from immune cell recognition and from the physical and chemical stresses of the blood circulation (Palumbo, Talmage et al. 2005). A hallmark of malignancy is the cancer cell’s ability to inhibit apoptosis. Platelets have been shown to reduce cancer cell apoptosis and improve cancer cell survival. For example, leukemia cells that underwent induced-apoptosis showed higher survival rates when incubated with platelets (Velez, Enciso et al. 2014). As the tumor grows, the needs for nutrients from the blood circulation increases. To sustain the metabolic requirement for the tumor growth, tumor cells are required to recruit additional blood vessels through angiogenesis. Platelet α-granules are rich in a wide variety of angiogenic factors including vascular endothelial growth factor (VEGF) and platelet derived growth factor (PDGF). Upon recruitment and activation by tumor cells, the release of these angiogenic molecules promotes the recruitment of blood vessels at the tumor site (Janowska- Wieczorek, Wysoczynski et al. 2005, Kuznetsov, Marsh et al. 2012). The anti-apoptotic properties of α-granule proteins such as VEGF and PDGF may also aid in the growth and survival of tumor cells (Le Gouill, Podar et al. 2004, Kosaka, Sudo et al. 2007, Tang, Arjunan et al. 2010, Vantler, Karikkineth et al. 2010). Consistent with the notion that platelet granules contribute to cancer cell development, platelet derived microvesicles in blood are present at a significantly higher level in cancer patients (Kim, Song et al. 2003, Koiou, Tziomalos et al. 2011, Peterson, Zurakowski et al. 2012).

Evidently, platelets play important roles in the pathology of inflammation, thrombosis, atherothrombosis, cancer and angiogenesis. The diverse assortment of molecules contained within platelet granules offer platelets the ability to regulate numerous critical biological process. While platelets are essential cells, dysregulation of platelet function can lead to dire consequences. Since platelet function involves the release of their granule proteins, understanding the nature of platelet secretory granules is imperative to the development of treatment strategies to modulate normal and abnormal platelet function.

1.2 Megakaryocytes and platelet biogenesis

Platelets are produced from megakaryocytes (MKs) via a unique developmental process. Representing less than one percent of the bone marrow population, MKs are also found in the lung and spleen, although it is unclear how much MKs residing outside the marrow contribute to platelet production. MKs are derived from pluripotent hematopoietic stem cells (HSCs) identified by the expression of CD34 (Ogawa 1993, Deutsch and Tomer 2013). Megakaryopoiesis (the differentiation of HSCs into MKs) and thrombopoiesis (the production of platelets) are regulated by the cytokine thrombopoietin (TPO) (Debili, Wendling et al. 1995, Kaushansky, Broudy et al. 1995), which is secreted by hepatocytes and bound via cell surface c-Mpl receptors expressed by circulating platelets, MKs and their HSC progenitors. When the circulating platelet concentration is high, little TPO enters the bone marrow and thrombopoiesis is minimal, but if the platelet count drops thrombopoiesis is stimulated (Jelkmann 2001, Scheding, Bergmann et al. 2002). TPO is required for HSC commitment to the MK lineage, and it supports but is not essential for the survival and maturation of MKs (Debili, Wendling et al. 1995, Kaushansky, Broudy et al. 1995, Matthews, Thevenin et al. 2011). There may also be a TPO-independent mode of MK development, but its role in normal thrombopoiesis has not been determined.

Mature MKs on average produce thousands of platelets each (Kaufman, Airo et al. 1965). In order to generate the required quantities of platelet proteins, organelles and membranes, maturing MKs undergo a massive increase in size as they pass through several rounds of endomitosis, where nuclear DNA is replicated but cell division does not occur (Patel, Hartwig et al. 2005, Geddis, Fox et al. 2007, Lordier, Jalil et al. 2008). Mature human MKs can attain diameters >100 μm and can have nuclear DNA equivalent up to 128N (Machlus, Thon et al. 2014), with proportionate increases in the ability to synthesize proteins, membranes, vesicles,

15 microtubules and other cellular components. As maturation proceeds, α-granule cargo (e.g. VWF) is loaded (Heijnen, Debili et al. 1998, Italiano and Battinelli 2009) and MKs develop an extensive invaginated membrane system (IMS), also known as the demarcation membrane system (DMS). The DMS serves as a plasma membrane reservoir for the final stage of platelet biogenesis (Schulze, Korpal et al. 2006, Eckly, Heijnen et al. 2014), when MKs extend long cytoplasmic protrusions known as proplatelets into bone marrow sinusoidal blood vessels (Becker and De Bruyn 1976, Junt, Schulze et al. 2007). Proplatelet elongation is driven by the extension of microtubules via polymerization and the sliding of tubules within bundles with the aid of dynein motors (Patel, Richardson et al. 2005, Bender, Thon et al. 2015). The microtubules loop around at the proplatelet tip before travelling backwards, forming a teardrop-shaped swelling known as a proplatelet bud (Italiano, Lecine et al. 1999).

Bending of proplatelets also occurs under the influence of actin polymerization forces, which creates loops or swellings along the proplatelet shaft that can serve as branching points for the generation of new extensions and sites of platelet assembly (Italiano, Lecine et al. 1999, Rojnuckarin and Kaushansky 2001). Proteins and organelles are transported along proplatelet microtubule tracks by kinesin motors from the MK cell body towards nascent platelets (Richardson, Shivdasani et al. 2005, Italiano, Patel-Hett et al. 2007) that resemble beads on a string. As proplatelets extend into sinusoidal blood vessels, the shear forces of blood flow release nascent platelets into circulation (Italiano, Lecine et al. 1999, Thon, Montalvo et al. 2010) while the MK cell body shrinks and the nucleus compresses. Eventually all that remains is a vestigial MK nucleus surrounded by thin layer of cytoplasm that is cleared by macrophages (Morison, Cramer Borde et al. 2008, Machlus, Thon et al. 2014). A flow sequence of MK development and maturation is illustrated in Fig 1.5 and the visualization of a proplatelet structure is shown in Fig. 1.6.

Many aspects of the cellular mechanisms involved in megakaryopoiesis and thrombopoiesis remain to be elucidated. Progress has been limited by the scarcity of MKs within the bone marrow population and the lack of suitable cell culture models. Several transcription factors have been linked to megakaryopoiesis, with the more well-characterized being GATA1 and friend of GATA1 (FOG1), both implicated as major regulators of MK maturation (Crispino 2005). Knockout of GATA1 in mice results in thrombocytopenia and the increased presence of small MKs within the bone marrow that have decreased ploidy and expression of granule cargo proteins, and immature DMS (Shivdasani, Fujiwara et al. 1997, Takahashi, Komeno et

16 al. 1998). FOG1 interacts with GATA1 to stimulate GATA1 activity, and this interaction has been shown to be essential for the differentiation of MKs and erythrocytes. Defects in GATA1-FOG1 interaction lead to thrombocytopenia, reduced MK differentiation and presence of immature MKs (Tsang, Visvader et al. 1997, Chang, Cantor et al. 2002).

Figure 1.5. Megakaryocyte development and maturation. Murine bone marrow MKs are grown in culture and imaged with immunofluorescence confocal microscopy. MKs are stained for DNA with DAPI (cyan), and immunostained for VWF (red), P-selectin (green) and α-tubulin (violet). The sequences represent the development and maturation of MKs up until proplatelet production. MKs undergo endomitosis (visualized by the rearrangement of microtubules at the nucleus) to achieve polyplodization and cell size expansion. The formation of long cytoplasmic extensions from the MK surface signifies the initiation of proplatelet production. During this process, MK proteins and organelles such as α-granules, as dictated by VWF and P-selectin staining, are transported along the proplatelet shaft towards the proplatelet tip. As the cytoplasm transforms into growing proplatelets, the MK cell body condenses, leaving a rimmed nucleus. The nucleus is eventually excised from the proplatelet structure. Scale bar = 20 µm.

Figure 1.6. Visualization of proplatelet structure. Mouse MKs are grown in culture and imaged via differential interference contrast (DIC) microscopy. Examples of some of the features of proplatelets are marked, including proplatelet swelling, tip, branch point and shaft. Each platelet-sized swelling at the proplatelet tip resembles a nascent platelet. Scale bar= 5 μm. Figure adapted and reprinted with permission (Patel, Hartwig et al. 2005).

1.3 Platelet structures and secretory granules 1.3.1 Overview

The visualization of platelet ultrastructure enables the characterization of platelets. Platelets are lined with circumferential microtubule bundles that form into bands of coiled rings. These microtubule coils confer integrity to the cytoskeleton and define the discoid shape of platelets (White and Burris 1984). Platelets possess extensive internal membrane systems known as the open canalicular system (OCS) and dense tubular system (DTS). OCS and DTS are not interconnected, and while OCS is continuous with the plasma membrane the DTS is not. Akin to the DMS in MKs, the OCS is a complex network of surface tunnels that are formed from the invagination of the plasma membrane (Selvadurai and Hamilton 2018). The OCS

18 provides additional surfaces for the endocytosis of plasma proteins and the externalization of secretory granules (Flaumenhaft 2003), and it also serves as a source of plasma membrane for platelet shape change and spreading following activation (Grouse, Rao et al. 1990). Derived from the MK smooth endoplasmic reticulum (ER), the DTS consists of thin tubules and functions in the regulation of platelet activation via the storage and release of calcium (Ebbeling, Robertson et al. 1992).

Three major classes of secretory granules have been identified in platelets: α-granules, δ- granules and lysosomes, each containing a unique set of cargo molecules (Blair and Flaumenhaft 2009, Heijnen and van der Sluijs 2015). α-granules are the most abundant, followed by δ-granules and lysosomes. Upon activation these granules fuse with the plasma membrane or OCS to release their contents (Flaumenhaft 2003), and this fusion also supplies additional membrane for the expansion of activated platelets (Berger, Masse et al. 1996, White 1999). The anatomy and some of the hallmark features of platelets are shown in Fig 1.7.

Figure 1.7. Anatomy of a platelet. Representative ultrastructural image of a platelet from a normal donor. Some of the hallmark features of a platelet are indicated. AG: α-granules, DB: δ-granule, OCS: open canalicular system, M: mitochondria, MT: microtubule coils. Scale bar = 500 nm. TEM performed by Hilary Christensen.

1.3.2 δ-granule biogenesis in megakaryocytes and platelets

1.3.2.1 Overview

δ-granules have been categorized as lysosome-related organelles (LRO), which as a group show similarities in biogenesis, yet have distinct aspects of structure, function and contents (Huizing, Helip-Wooley et al. 2008). Other LROs include melanosomes and natural killer cell lytic granules, which function in pigmentation and cytotoxicity respectively (Dell'Angelica, Mullins et al. 2000). δ-granules average 200 nm in diameter with a frequency of 3-8 granules per platelet (Ruiz, Lea et al. 2004, Urban, Pluthero et al. 2017). In ultrastructural imaging, δ- granules appear spherical with an electron dense central core (Saultier, Vidal et al. 2017). Constituents of δ-granules are primarily small molecules and ions, including adenosine diphosphate (ADP), adenosine triphosphate (ATP), potassium, polyphosphates, calcium and serotonin. The high concentration of ions and molecules gives the δ-granule core its dark appearance in TEM (McNicol and Israels 1999, Ruiz, Lea et al. 2004, Fitch-Tewfik and Flaumenhaft 2013). Similar to the acidic pH observed in lysosomes and other LRO, the intraluminal pH of δ-granules is maintained at a pH of 5.4 by membrane hydrogen ion pumps (Dean, Fishkes et al. 1984, Huizing, Helip-Wooley et al. 2008). δ-granules can be identified via the membrane proteins CD63 and LAMP-1, which they share with lysosomes (McNicol and Israels 1999, Knight, Gomez et al. 2017). The uniqueness of LROs is indicated by the observation that genetic defects can specifically affect one type but not the others (Raposo, Marks et al. 2007, Huizing, Helip-Wooley et al. 2008). Our understanding of δ-granule biogenesis is largely based on studies of protein defects associated with δ-granule deficiencies, including those causing Hermansky-Pudlak syndrome (HPS), Chediak-Higashi syndrome (CHS) and Griscelli syndrome (GS).

1.3.2.2 Platelet δ-granule deficiencies 1.3.2.2.1 Hermansky-Pudlak syndrome

Hermansky-Pudlak syndrome (HPS) was first described in 1959 as an autosomal recessive disease characterized by oculocutaneous albinism and hemorrhagic diathesis despite normal platelet counts (Hermansky and Pudlak 1959). Later studies linked oculocutaneous albinism to defects in the formation of melanosomes, and bleeding diathesis to platelet δ-granule deficiencies (Gahl, Brantly et al. 1998, Huizing, Helip-Wooley et al. 2008). Other clinical symptoms may include granulomatous colitis, immune deficiency and pulmonary fibrosis. The manifestation and severity of HPS symptoms are variable among patients, where several

20 subtypes are evident (Huizing, Helip-Wooley et al. 2008). Analysis of HPS platelet ultrastructure revealed the absence of δ-granules but normal α-granules, suggesting distinct biogenesis pathways of these secretory granules (Huizing, Parkes et al. 2007). To date, ten HPS subtypes have been reported in humans: AP3B1, AP3D1, BLOC1S3, BLOC1S6, DTNBP1, HPS1, HPS3, HPS4, HPS5 and HPS6. Each HPS subtype combines to form one of the four complexes with known roles in vesicular trafficking: three subtypes of biogenesis of lysosome-related organelles complex (BLOC-1, BLOC-2 and BLOC-3) and adaptor protein complex 3 (AP-3). Specifically, DTNBP1, BLOC1S3 and BLOC1S6 are constituents of BLOC- 1 complex; HPS3, HPS5 and HPS6 form BLOC-2 complex; HPS1 and HPS4 form BLOC-3 complex; AP3B1 and AP3D1 are subunits of AP-3 complex (Gahl, Brantly et al. 1998, Morgan, Pasha et al. 2006, Huizing, Helip-Wooley et al. 2008, Badolato, Prandini et al. 2012, Ammann, Schulz et al. 2016).

Interestingly, BLOC-1, BLOC-2 and AP-3 complexes have the capacity to interact with each other. AP-3 is able to interact with BLOC-1, BLOC-1 is able to interact with BLOC-2, though AP-3 is unable to interact with BLOC-2 (Di Pietro, Falcon-Perez et al. 2006). While all three complexes localize to early endosomal tubules, AP-3 acts first by organizing the formation of clathrin vesicle budding, followed by the actions of BLOC-1 to direct the vesicles to endosomal intermediates, and finally BLOC-2 targets cargo to LROs (Dell'Angelica, Klumperman et al. 1998, Di Pietro, Falcon-Perez et al. 2006, Salazar, Craige et al. 2006, Setty, Tenza et al. 2007, Dennis, Mantegazza et al. 2015). Deficiencies in the combination of BLOC-1, BLOC-2 and AP-3 can have different phenotypes in mice. Mice with double knockout of BLOC-1 and BLOC-2 are indistinguishable in coat color from mice with single knockout of BLOC-1, mice with dual knockout of BLOC-1 or BLOC-2 with AP-3 show increased coat color deficiency relative to mice with single knockout of BLOC-1, BLOC-2 or AP-3 (Di Pietro, Falcon-Perez et al. 2006). These data suggest that BLOC-1 and BLOC-2 act in the same molecular pathway, while AP-3 can be involved in sorting pathways that are independent of BLOC-1/2. This hypothesis was corroborated by a subsequent study where it was shown that AP-3 and BLOC-1/2 can independently mediate different LRO sorting pathways to selectively sort melanosome cargo proteins (Setty, Tenza et al. 2007).

Divergent from the other HPS complexes, BLOC-3 acts independently of AP-3 and does not interact with AP-3, BLOC-1 or BLOC-2 (Martina, Moriyama et al. 2003, Di Pietro, Falcon- Perez et al. 2006). Relative to defects with other HPS subtypes, HPS patients with BLOC-3

21 deficiencies are often associated with more severe forms of oculocutaneous albinism, hemorrhagic diathesis, granulomatous colitis, and the often lethal disorder pulmonary fibrosis (Gahl, Brantly et al. 2002, Hussain, Quezado et al. 2006, Huizing, Helip-Wooley et al. 2008). BLOC-3 functions as a guanine nucleotide exchange factor (GEF) for RAB32 and RAB38, and since BLOC-3 binds to GTP-bound RAB9 it may also function as an effector for RAB9 (Kloer, Rojas et al. 2010, Gerondopoulos, Langemeyer et al. 2012). RAB9, RAB32 and RAB38 have all been implicated in the delivery of cargo to LROs (Wasmeier, Romao et al. 2006, Mahanty, Ravichandran et al. 2016). As mice with double knockout of BLOC-1 and BLOC-3 are indistinguishable in coat color from mice with single knockout of BLOC-1, it suggest that BLOC-1/3 may act in the same molecular pathway (Nazarian, Falcon-Perez et al. 2003). Later study has shown that BLOC-3 can also selectively sort melanosome cargo in a pathway independent of BLOC-1/2, suggesting that BLOC-3 may participate in multiple sorting pathways (Bultema, Ambrosio et al. 2012). The function of BLOC-3 is likely mediated by RAB32/38 (Bultema, Ambrosio et al. 2012, Gerondopoulos, Langemeyer et al. 2012). RAB32/38 were also found to interact with BLOC-2, AP-3 and AP-1, and through these interactions mediate LRO protein trafficking from the early endosomes (Bultema, Ambrosio et al. 2012). Intriguingly, studies have recently revealed the requirement of BLOC-1 and BLOC- 3 in regulating the cycling of VAMP7, a v-SNARE required for melanogenesis, between endosomes and melanosomes. Specifically, VAMP7 and BLOC-1 mediate the delivery of cargo to melanosomes, while BLOC-3 regulates the recycling of VAMP7 from the melanosomes, a step that also involves the action of RAB32/38 (Dennis, Delevoye et al. 2016).

Studies of buff mice revealed the role of Class C VPS, also known as the HOPS complex, in δ-granule biogenesis. buff mice display features of HPS including hypopigmentation, bleeding diathesis and platelet δ-granule defects. The underlying cause is a single point mutation in vacuolar protein sorting 33A (VPS33A), a subunit of the homotypic fusion and protein sorting (HOPS) complex. Lysosomal functions are not affected in buff mice, suggesting that the action of HOPS complex is specific to melanosomes and platelets δ-granules. Because the defects observed in buff mice are mild, the potential of residual VPS33A activity should be considered (Suzuki, Oiso et al. 2003). The HOPS complex is a heterohexamer consisting of VPS33A, VPS16A, VPS41, VPS18, VPS11 and VPS39. Within this complex VPS33A binds to VPS16A to form a sub-complex that can interact with SNARES (Balderhaar and Ungermann 2013, Solinger and Spang 2013). HOPS functions as an effector of RAB7 and

22 tethers RAB7 late endosomes to LRO membranes. Subsequently, HOPS interacts with soluble NSF attachment protein receptors (SNAREs) present on the late endosome and on the LRO, thereby promoting SNARE complex assembly and facilitates membrane fusion (Caplan, Hartnell et al. 2001, Kim, Kramer et al. 2001, Poupon, Stewart et al. 2003, Richardson, Winistorfer et al. 2004). HOPS can also interact with AP-3 complex to facilitate the transport of cargo from the Golgi complex to vacuoles in yeast (Angers and Merz 2009). Defects in members of the HOPS complex are often lethal, as observed in Drosophila where larvae harboring dVps16A or dVps33A knockout fail to thrive. Conditional knockout of dVps16A or dVps33A in Drosophila eye revealed that both dVps16A and dVps33A are required for retinal pigmentation, autophagosome maturation and lysosomal trafficking (Pulipparacharuvil, Akbar et al. 2005, Akbar, Ray et al. 2009). Many of these protein subunits are unique in function and are not substitutable by their homologs. For example, lysosomal trafficking defects caused by the loss-of-function of dVps33A in Drosophila eye is not rescued by the transgenic expression of its homologue, dVps33B, thereby suggesting non-redundant functions (Akbar, Ray et al. 2009).

1.3.2.2.2 Chediak-Higashi syndrome

Chediak-Higashi syndrome (CHS) is an autosomal recessive disease caused by mutations in the lysosomal trafficking regulator (LYST) gene. LYST belongs to a family of proteins containing the beige and Chediak-Higashi (BEACH) domain (Burgess, Mornon et al. 2009). Clinical manifestations of CHS include hypopigmentation, neurologic dysfunction, bleeding diathesis and immunologic defects leading to chronic infections (Introne, Boissy et al. 1999, Huizing, Helip-Wooley et al. 2008, Kaplan, De Domenico et al. 2008). The hallmark of CHS is the presence of giant lysosomal granules in leukocytes due to dysregulated homotypic fusion of lysosomes (White and Clawson 1980, Irimajiri, Iwamoto et al. 1992, Jones, Stewart et al. 1992). Immunodeficiency in CHS is a combined result of neutropenia, impaired leukocyte chemotaxis and enlarged lysosomes in leukocytes failing to degranulate to form phagolysosomes, thus preventing the elimination of pathogens via phagocytosis (Clark and Kimball 1971, Renshaw, Davis et al. 1974, White and Clawson 1980). Further, CHS natural killer cells displayed reduced cytotoxicity due to inability to exocytose lytic granules via plasma membrane fusion (Gil-Krzewska, Wood et al. 2016). To date, despite the clear implications of LYST in regulating lysosomal size and functions, the precise role of LYST remains unclear and controversial. While initially LYST was proposed to function in the

23 prevention of lysosomal homotypic fusion (Tanabe, Cui et al. 2000, Tchernev, Mansfield et al. 2002, Mohlig, Mathieu et al. 2007), recent studies and analysis supported the role of LYST in promoting lysosomal fission (Durchfort, Verhoef et al. 2012, Cullinane, Schaffer et al. 2013).

Susceptibility to bleed in CHS is caused by platelet function impairments due to the absence or reduced platelet δ-granules despite normal α-granules and normal platelet counts (Rendu, Breton-Gorius et al. 1983, Introne, Boissy et al. 1999). Despite the absence or reduction in platelet δ-granules, the δ-granules that were observed were normal in size. CHS platelets display aggregation defects, have an increased ATP/ADP ratio, and are defective in serotonin uptake and release (Parmley, Poon et al. 1979, Introne, Boissy et al. 1999, Gunay-Aygun, Huizing et al. 2004). Similar to leukocytes, though with heterogeneity, giant granules of lysosomal nature are occasionally observed in CHS platelets (Parmley, Poon et al. 1979). In CHS melanocytes, melanosomes appeared enlarged and melanosome proteins were mislocalized (Introne, Boissy et al. 1999, Westbroek, Adams et al. 2007). Similar mis-sorting of lysosomal elastase and cathepsin G were observed in CHS neutrophils (Takeuchi, Wood et al. 1986). Analysis of perforin-containing granules in CHS natural killer cells reveals a similar trend of aberrant compartment protein trafficking (Gil-Krzewska, Wood et al. 2016). Yeast-two-hybrid screens using truncated LYST domains identified that LYST interacts with HRS, a SNARE complex implicated in the vesicular trafficking of ubiquitinated proteins from late endosomes to lysosomes (Tchernev, Mansfield et al. 2002, Raiborg, Rusten et al. 2003). Based on these findings, it is also likely that LYST functions in regulating the vesicular trafficking of lysosomal and LRO proteins.

1.3.2.2.3 Griscelli syndrome

Griscelli syndrome (GS) is a rare autosomal recessive disease characterized by defects in MYO5A, RAB27A and melanophilin (Mancini, Chan et al. 1998, Menasche, Pastural et al. 2000, Menasche, Ho et al. 2003). GS manifests in hypopigmentation and by primary neurological impairments when MYO5A is affected and by immunodeficiency and lymphohistiocytosis when RAB27A is implicated (Pastural, Barrat et al. 1997, Menasche, Pastural et al. 2000, Menasche, Ho et al. 2003). In melanocytes, MYO5A, RAB27A and melanophilin forms a complex that works in tandem to capture melanosomes at the cell periphery following their transport from the cell body, a step that is critical for the transfer of melanosomes to keratinocytes for pigmentation. Mutation of any member of the complex

24 results the perinuclear accumulation and clumping of melanosomes (Wu, Rao et al. 2002, Westbroek, Lambert et al. 2003, Van Gele, Dynoodt et al. 2009).

GS patients do not have an overt bleeding diathesis, hence platelet δ-granules have not been carefully examined. It is therefore uncertain whether GS implicates δ-granules based on the clinical cases. Initially, a RAB27A deficient mice, ashen, was reported to display significant bleeding disorders caused by the reduction of platelet δ-granules (Wilson, Yip et al. 2000). Ultrastructural analysis of ashen platelets revealed no additional abnormalities outside of δ- granule defects, though it was noted that the platelet and δ-granule defects seen in ashen are highly dependent on the genetic background of the mice (Novak, Gautam et al. 2002). In a subsequent study conflicting evidence was again observed with the ashen strain, where mice displayed no bleeding tendencies and had normal platelet δ-granule counts (Barral, Ramalho et al. 2002). Later evidence indicated that δ-granule deficiency in the original study was caused by the mutation of another gene that was also present in the mice, Slc35d3, rather than by the proposed Rab27A (Chintala, Tan et al. 2007). As no defects of other LRO were observed in SLC35D3 deficient mice, SLC35D3 appears to regulate δ-granule biogenesis specifically (Chintala, Tan et al. 2007). Studies of SLC35D3 in MKs revealed that while SLC35D3 localizes to the early endosomes, it does not localize to δ-granules. However, since the activity of SLC35D3, BLOC-1 and AP-3 are inter-dependent, it is postulated that SLC35D3, like BLOC-1 and AP-3 in mediating LRO formation, functions in mediating δ- granule biogenesis from the early endosomes (Meng, Wang et al. 2012). Based on these findings and despite SLC35D3 not being directly implicated in GS, it is apparent that SLC35D3 partakes an important role in δ-granule biogenesis. However, it is unclear whether SLC35D3 directly regulates the formation of δ-granules or acts as an intermediary during this process.

Considering the evidence that RAB27A-deficient ashen ultimately did not exhibit platelet δ- granule defects and the lack of bleeding tendencies in GS patients, it seems unlikely that RAB27A partakes in δ-granule biogenesis. Though interestingly, it was shown that RAB27A and its isoform, RAB27B, both localize primarily to δ-granules in platelets, suggesting their potential roles in δ-granule function (Barral, Ramalho et al. 2002). The functions of RAB27A and RAB27B are at least partially redundant as the transgenic expression of RAB27B can rescue the hypopigmentation phenotype in RAB27A-deficient ashen mice (Barral, Ramalho et al. 2002). In melanocytes, RAB27A performs the function of docking melanosomes to the cell

25 periphery by binding to melanophilin-MYO5A complex that are present at the cortical actin network (Wu, Rao et al. 2002, Westbroek, Lambert et al. 2003). This corroborates the studies of RAB27A and RAB27B in Hela cells where RAB27B functions to transport and retain exosomes at the cell periphery, allowing RAB27A to dock the exosomes to the plasma membrane (Ostrowski, Carmo et al. 2010). These findings suggest that RAB27A and RAB27B also have non-redundant functions.

Furthermore, as opposed to RAB27A being ubiquitously expressed, the expression of RAB27B is restricted to a few tissue and cell types including platelets (Barral, Ramalho et al. 2002, Zhao, Torii et al. 2002, Chen, Guo et al. 2003, Chen, Li et al. 2004). Considering that RAB27B is selectively expressed in platelets and that RAB27B localizes to δ-granules, it raises the likelihood that RAB27B may play a critical role in platelet function. Thus, despite RAB27A showing no phenotypic changes upon deletion, RAB27B-deficient mice were generated in search of a role in platelet function. RAB27B-deficient mice showed platelet function impairments as a result of decreased δ-granule counts, whereas the deletion of RAB27A has no distinctive effects on platelet ultrastructure and functions (Tolmachova, Abrink et al. 2007). Consistent with the role of RAB27B in exosome secretion, the secretion of δ-granules is decreased upon RAB27B knockout whereas α-granule externalization is unaffected (Tolmachova, Abrink et al. 2007). The reduced secretion of δ-granules appears to correlate linearly with the reduced numbers of δ-granules present in the cell, however the precise role of RAB27B in platelet δ-granule secretion requires further studies. Nonetheless, it is proposed that RAB27B is required for δ-granule biogenesis and may also participate in the secretion of δ-granules.

1.3.2.3 Model of δ-granule biogenesis

Through the studies of human diseases characterized by LRO and platelet δ-granule deficiencies, much has been learnt regarding the events and mechanisms involved in δ- granule formation. δ-granule cargo sorting, as with other LRO, begins at the early endosomes where the AP-3 complex directs the formation of clathrin vesicle budding. These vesicles are then directed to late endosomes or targeted for δ-granules. Along the route where the vesicles are targeted for δ-granules from the early endosomes, BLOC-1 directs the exit of vesicles from the early endosomes into endosomal intermediates and BLOC-2 targets the final delivery to the maturing δ-granules. SLC35D3 cooperates with AP-1 and BLOC-1 in delivering δ-granule cargoes from the early endosome. BLOC-3 can act independently of AP-

3 and BLOC-1/2 and selectively delivers cargoes to maturing δ-granules. There also exist distinct sorting pathways for different δ-granule cargoes that are regulated independently by AP-3 or BLOC-1/2 and do not require the cooperation between AP-3 and BLOC-1/2. VAMP7 SNARE mediates the fusion of early endosomes, containing δ-granule cargoes, to maturing δ-granules and is a step that is mediated by BLOC-1. Upon cargo delivery, VAMP7 fuses with the target membranes and its retrieval from the δ-granules back towards the early endosomes is mediated by BLOC-3. In the route where δ-granule cargoes are targeted to the late endosomes, the HOPS complex regulates the delivery of late endosomal cargoes into nascent δ-granules. LYST/CHS is proposed to inhibit fusion or promote fission of LRO where defects in LYST leads to enlarged lysosomes and melanosomes. This however does not explain the absence of δ-granule in CHS platelets, suggesting an alternative function of LYST in platelet δ-granule biogenesis. As studies have suggested that LYST defects also lead to the missorting of cargoes in multiple cell types that harbor LRO, it is conceivable that LYST also functions in the vesicular trafficking of δ-granule cargoes in platelets. RAB27B localizes to δ-granules, regulating its formation and secretion.

1.3.3 α-granule biogenesis in megakaryocytes and platelets 1.3.3.1 Overview

α-granules are the most abundant platelet secretory vesicles, averaging 50 to 80 per cell (Chen, Lo et al. 2017). In contrast to δ-granules, which contain mostly small molecules and ions, α-granules primarily carry proteins and account for the majority of proteins secreted from platelets during activation (Blair and Flaumenhaft 2009). These proteins act in a variety of cellular processes. Adhesive proteins such as thrombospondin-1 (TSP1), VWF and FGN, proteases such as prothrombin and clotting factors such as factor V are all key elements of primary and secondary hemostasis. PDGF and VEGF enhance wound healing and promote angiogenesis. NAP-2, PF4 and the membrane protein P-selectin facilitate immune responses (McNicol and Israels 2008, Wolberg and Campbell 2008, Blair and Flaumenhaft 2009, Smyth, McEver et al. 2009, Peterson, Zurakowski et al. 2012).

α-granule regulation is under active investigation. It has been proposed that specific subsets of α-granules differentially release their contents depending on the activation agonist (Italiano, Richardson et al. 2008), but this has been challenged by the proposition that proteins are released according to their orientation within the granule relative to the fusion pore at rates determined by the kinetics of exocytosis (Kamykowski, Carlton et al. 2011). α-granules

27 display considerable morphological heterogeneity. Ultrastructural studies show that most are 200-500 nm spherical structures, but tubular and multivesicular subtypes are also present. Immunogold microscopy studies indicate that different α-granule subtypes may differ in their contents. For example, VWF and P-selectin are primarily found in spherical α-granules, whereas the presence of FGN is equally prominent in all types (van Nispen tot Pannerden, de Haas et al. 2010). Whether different structures indicate different functional classes of α- granules remains to be determined, as do many aspects of α-granule biogenesis that are poorly understood. Recent progress in this area has come from studies of two hereditary α- granule deficiencies: arthrogryposis, renal dysfunction and cholestasis (ARC) syndrome and gray platelet syndrome (GPS).

1.3.3.2 α-granule trafficking 1.3.3.2.1 Trafficking of α-granule proteins

Most proteins carried by platelet α-granules are synthesized by MKs, but some are endocytosed by MKs or platelets. Platelets are also capable of synthesizing α-granule proteins such as PAI-1 (Brogren, Karlsson et al. 2004). This raises an important question regarding how platelets regulate the synthesis and trafficking of these secretory proteins. Conventionally, secretory proteins are first processed in the Golgi apparatus prior to secretion or delivery into specialized organelles, as is the case of VWF synthesis in MKs (Schick, Walker et al. 1997, Farhan and Rabouille 2011). Structured Golgi complex is however not seen in platelets (Yadav, Williamson et al. 2017). Though recent evidence has suggested an alternative unconventional secretion pathway that bypasses the Golgi apparatus, it is unclear whether platelets utilize similar mechanisms (Rabouille 2017). Further, the differences in structures and organelle compositions in MKs and platelets may suggest that different mechanisms of regulation exist for α-granule protein trafficking. This raises the need to differentially identify the mechanism of α-granule protein trafficking in MKs and in platelets.

The abundant α-granule protein FGN is not synthesized by MKs (Handagama, Rappolee et al. 1990, Louache, Debili et al. 1991). Endocytosis of FGN into MKs and platelets is mediated by integrin αIIbβ3 surface receptors and requires the functions of ADP-ribosylation factor 6 (ARF6) in platelets (Handagama, Bainton et al. 1993, Handagama, Scarborough et al. 1993, Huang, Joshi et al. 2016). MK-synthesized proteins such as VWF and TSP1 undergo the classical secretory pathway for trafficking into α-granules (Majack, Cook et al. 1985, Peyvandi, Garagiola et al. 2011). VWF is first translated as pre-pro-VWF, where a signal peptide and a

28 pro-peptide are present at the N-terminal. In the ER the signal peptide is cleaved, generating pro-VWF, followed by glycosylation and the formation of pro-VWF dimers via C-terminal disulfide linkage. Within the Golgi apparatus, additional glycosylation and sugar processing occurs, followed by the cleavage of pro-peptide from pro-VWF dimers, producing mature VWF dimers. Mediated by the cleaved pro-peptides, mature VWF dimers further multimerize with other dimers via N-terminal disulfide linkages. These VWF multimers are then transported into early endosomes or directly into nascent α-granules (Schick, Walker et al. 1997, Sadler 2009, Peyvandi, Garagiola et al. 2011). In ultrastructural studies, VWF multimers in α-granules appear as distinct tubular assemblies that are 12 nm in length (Cramer, Meyer et al. 1985, van Nispen tot Pannerden, de Haas et al. 2010).

Few studies have examined the specifics of α-granule biogenesis, and current knowledge of cargo protein trafficking comes from a single ultrastructural study reported in 1998 where Heijnen et al. examined and traced the movements of endocytosed and biosynthesized α- granule proteins across MK organelles via immunogold electron microscopy. Tracking of tracer proteins in MKs revealed that following endocytosis these proteins are first detected near the plasma membrane in tubulo-vesicular endosomes which may resemble early endosomes. The proteins are then transported to multivesicular bodies I (MVB I), late endosomes with numerous intraluminal vesicles which mature into multivesicular bodies II (MVB II), late endosomes with intraluminal vesicles and electron dense matrices, then finally to α-granules. The close association of tracer proteins with the electron-dense matrices of MVB II suggests that α-granules may be derived from these structures. Similarly, biosynthesized α-granule proteins are trafficked along a similar pathway where P-selectin and VWF are found in transport vesicles, Golgi complex, MVB I, MVB II and α-granules (Heijnen, Debili et al. 1998). The precise details and mechanisms of α-granule biogenesis and α- granule protein trafficking are still largely unknown.

1.3.3.2.2 Endosome maturation

α-granule biogenesis involves the trafficking of cargo proteins to nascent α-granules (Heijnen, Debili et al. 1998), where multiple routes have been proposed. The convergence of VWF and FGN in late endosomal MVBs strongly suggests that the endocytic cycle is a prominent pathway of α-granule protein trafficking (Heijnen, Debili et al. 1998). This pathway describes a series of trafficking events that begin at early endosomes (EE) for both synthesized and endocytosed cargo proteins. EE are phosphatidylinositol-3-phosphates (PI3P) rich tubular-

29 vesicular organelles that act as the central sorting platforms within the endocytic cycle. Proteins derived from both the plasma membrane via RAB5 GTPase activity, and trans-Golgi network (TGN) via AP-1 adaptor complex regulation, accumulate within EE for directed sorting into specialized organelles, including LRO and δ-granules (Dell'Angelica, Mullins et al. 2000, Jovic, Sharma et al. 2010, Luzio, Hackmann et al. 2014, Marat and Haucke 2016). Alternatively, EE mature in MVBs, late endosomal organelles characterized by the presence of intraluminal vesicles (ILV), through a process that requires the action of endosome sorting complex required for transport (ESCRT) complexes. ESCRT complexes direct the inward- budding and fission of the limiting membranes of EE to generate ILV, which serve to selectively compartmentalize membrane proteins (Piper and Katzmann 2007). Marked by the transient switching of markers including RAB5 to RAB7 GTPase and PI3P to phosphatidylinositol 3,5-bisphosphate (PI(3,5)P2), the endosomal structure matures as ILV accumulate and eventually give rise to specialized cellular organelles such as α-granules, δ- granules and LRO, or undergo degradation through lysosomal fusion via the formation of endolysosomes (Heijnen, Debili et al. 1998, Huotari and Helenius 2011, Luzio, Hackmann et al. 2014).

Endosomal maturation is thought to be facilitated by multiple factors, including the formation of membrane contact sites (MCS) between ER and endosomes. The non-fusogenic ligation between the membrane proteins of peripheral ER and target endosome establishes the basis of MCS, promoting lipid transfer and facilitating signaling events such as vesicular fusion and fission (Phillips and Voeltz 2016). Through high-resolution 3D electron microscopy, it has been revealed that endosomes are constantly in contact with the ER, and intriguingly the degree of MCS formation increases as the endosomes mature, suggesting a role for MCS in endosomal maturation (Friedman, Dibenedetto et al. 2013). The formation of membrane contacts between ER protein vesicle-associated membrane protein-associated protein A (VAP-A) and late endosome protein StAR-related lipid transfer domain protein 3 (STARD3) regulates the conformation and dynamics of late endosomes (Alpy, Rousseau et al. 2013). Similarly, the formation of a distinct ER-late endosome tether between ER protein VAP-A and late endosome protein oxysterol-binding protein-related protein-1L (ORP1L) regulates the transport and positioning of late endosomes, a process that is correlated with endosomal maturation (Rocha, Kuijl et al. 2009, Huotari and Helenius 2011). The interaction between VAP-A and ORP1L also prevents fusion of late endosomes by inhibiting the interaction of Rab-interacting lysosomal protein (RILP) with HOPS complex, which together normally

30 mediates the trafficking and fusion of late endosomes (van der Kant, Fish et al. 2013). Both RILP and HOPS complex have previously been implicated in endosomal maturation (Harrison, Brumell et al. 2004, Liang, Lee et al. 2008). Further, ER-endosome tethering has been shown to be essential for endosomal fission, a critical process of protein trafficking in the endocytic pathway (Rowland, Chitwood et al. 2014). While the discovery of MCS in endosomal processes is recent, emerging evidence supports the role of MCS in regulating endosome maturation, dynamics and functions. MCS extends beyond ER-endosome interactions. MCS between VAP-A on ER and ORP1L on autophagosomes regulates autophagosome maturation and its fusion with lysosomes (Wijdeven, Janssen et al. 2016). The ER-resident SNARE, SEC22B, shown to be involved in vesicular trafficking events and phagosome maturation, also forms MCS with plasma membrane protein syntaxin-1 to promote cell growth and plasma membrane expansion via lipid transfer (Cebrian, Visentin et al. 2011, Petkovic, Jemaiel et al. 2014).

1.3.3.3 Platelet α-granule deficiencies 1.3.3.3.1 Gray platelet syndrome

Gray platelet syndrome (GPS) is an autosomal recessive bleeding disorder caused by mutations in the gene encoding neurobeachin-like 2 (NBEAL2) (Albers, Cvejic et al. 2011, Gunay-Aygun, Falik-Zaccai et al. 2011, Kahr, Hinckley et al. 2011). In patients GPS causes a bleeding diathesis, macrothrombocytopenia, splenomegaly and progressive myelofibrosis (Nurden and Nurden 2007, Gunay-Aygun, Zivony-Elboum et al. 2010). Ultrastructural studies have shown the virtual absence of α-granules in GPS platelets, while δ-granule and lysosome numbers and structures are unaffected (Breton-Gorius, Vainchenker et al. 1981). Analysis of GPS platelets revealed platelet aggregation impairment and absence of platelet α-granule contents, while δ-granule and lysosome secretion and function appear unaffected (Gerrard, Phillips et al. 1980). Based on these data, bleeding diathesis in GPS is attributed to impairment of platelet function as a result of α-granule deficiency. It should be noted that signs of progressive myelofibrosis in GPS indicate a change in the bone marrow environment that may have resulted from causes other than platelet function defects. On the other hand, splenomegaly is often associated with myelofibrosis and is associated with extramedullary hematopoiesis (Randhawa, Ostojic et al. 2012)

GPS platelets are generally large, and as a result of α-granule loss appear gray when visualized with light microscopy, hence the name GPS (Nurden and Nurden 2007, Gunay-

Aygun, Zivony-Elboum et al. 2010). GPS platelets and MKs also appear highly vacuolated when compared to normal platelets and MKs (Fig.1.8A-B and Fig. 1.9). Interestingly, despite the absence of classical α-granules, P-selectin was detected along the limiting membrane of vacuole-like vesicles in GPS platelets. Further, it was shown that these vacuole-like vesicles are capable of fusion and secretion upon platelet activation, and despite the paucity of biosynthesized α-granule proteins, endocytosed albumin is detectable within these structures. From this evidence it was postulated that these vacuole-like vesicles may represent immature α-granules, or ghost granules, and that the inherent defect in GPS platelets involves the trafficking of α-granule proteins (Rosa, George et al. 1987). Mass spectrometry analysis of ghost α-granules from GPS platelets showed corroborating results. In this study, it was shown that biosynthesized α-granule proteins such as VWF and TSP1 are absent from GPS ghost granules, and interestingly the presence of endocytosed α-granule proteins such as FGN and albumin is less affected, albeit still decreased when compared to control platelets (Maynard, Heijnen et al. 2010).

Figure 1.8. Ultrastructure of platelets from ARC and GPS patients. TEM images of platelets from a healthy individual (A), GPS patient (B) and ARC patients with mutation in VPS16B (C) or VPS33B (D). GPS and ARC platelets are larger in size and display α-granule deficiency. In contrast, α-granules are common in normal platelets as indicated by the white arrows. Scale bar = 500 nm. Figure adapted and reprinted with permission (Chen, Lo et al. 2017)

Figure 1.9. Ultrastructure of GPS megakaryocytes. TEM image of a GPS bone marrow MK (A) and a normal bone marrow MK (B). GPS MK appears highly vacuolated, showing the absence of α- granules. This directly contrasts the normal MK where α-granules are commonly seen as indicated by the white arrows. Scale bars = 2 μm. Figure adapted and reprinted with permission (Chen, Lo et al. 2017).

It is apparent that α-granule proteins are not trafficked into α-granules in GPS platelets, however, the fate of these proteins is unclear. Ultrastructural examination provided some insights into the trafficking of α-granule proteins in GPS MKs. In immature MKs, it was shown that electron-dense granules of 50-100 nm in diameter were originally observed near the Golgi complex but dissipated over time as MK developed. These granules are thought to be α-granule precursors that failed to mature. At the same time, the appearance of electron- dense material near the MK DMS indicates aberrant externalization (Breton-Gorius, Vainchenker et al. 1981). Consistently, studies of VWF localization in GPS MKs have shown that VWF is deposited in the DMS following its appearance near the Golgi complex (Cramer, Vainchenker et al. 1985, Drouin, Favier et al. 2001). Altogether, these data indicate that the fundamental defect of α-granule biogenesis in GPS MKs and platelets is likely the aberrant trafficking of cargo proteins. These observations also raise the hypothesis that the aberrant release of α-granule cargo, which includes many growth factors, by MKs may contribute to the development of myelofibrosis and splenomegaly (Nurden and Nurden 2007) in GPS patients.

1.3.3.3.2 Arthrogryposis, renal dysfunction and cholestasis syndrome

Arthrogryposis, renal dysfunction and cholestasis (ARC) syndrome is a fatal autosomal recessive multisystem disorder caused by mutations in genes encoding VPS33B or VPS16B (Urban, Li et al. 2012). At birth, ARC syndrome manifests in functional defects of musculoskeletal system, kidney, liver and central nervous system. Prognosis is poor and patients generally do not survive past the first year of life as a result of complications with immunodeficiency, dehydration, acidosis and internal hemorrhages (Gissen, Tee et al. 2006). Bleeding diathesis in ARC syndrome ranges from mild to severe, manifesting in internal hemorrhages and instances of life-threatening bleeding episodes (Elmeery, Lanka et al. 2013, Saadah, Bokhari et al. 2013). Despite predisposition to bleeding, ARC patients generally have normal platelet counts. Under blood film visualization ARC platelets generally appear large and pale (Lo, Li et al. 2005, Gissen, Tee et al. 2006). Electron microscopy studies revealed the absence of α-granules (Fig. 1.8A,C-D) and the presence of δ-granules, while immunoblot analysis confirmed the absence of the α-granule membrane constituent P- selectin and granule cargo proteins such as VWF and FGN (Lo, Li et al. 2005, Urban, Li et al. 2012).

Loss of expression of either VPS33B or VPS16B causes ARC syndrome (Urban, Li et al. 2012). The role of VPS33B in α-granule biogenesis was examined in mice with conditional knockout of Vps33b in MKs, Vps33bfl/fl-ERT2, which recapitulate the ARC-associated bleeding diathesis and produce platelets lacking α-granules but show normal numbers of δ-granules, indicating that VPS33B specifically affects α-granule development. Analysis of Vps33bfl/fl-ERT2 MKs revealed that the loss of VPS33B delays MK maturation and reduces the numbers of mature MVB II. VWF production in Vps33bfl/fl-ERT2 MK is normal, but trafficking within the endocytic pathway is abnormal with VWF in MVB II being significantly reduced (Bem, Smith et al. 2015). Based on these data, it is proposed that VPS33B functions in the trafficking of α- granule proteins from TGN to MVB, which facilitates MVB maturation and promotes α-granule biogenesis. This model is consistent with localization studies of VPS33B and VPS16B where VPS33B is observed to reside with MVB and α-granules and VPS16B is found to localize with TGN, MVB and α-granules (Lo, Li et al. 2005).

1.3.3.4 Model of α-granule biogenesis

Fig. 1.10 illustrates a proposed model of α-granule biogenesis based on current knowledge. VPS33/16B is proposed to act from the trans-Golgi, mediating α-granule protein trafficking towards MVB I, and continues to facilitate the maturation of MVB I to MVB II. Since P-selectin is undetectable in ARC platelets while P-selectin vesicles, hypothesized as α-granule precursors, are observed in GPS platelets, it is postulated that VPS33B/16B is responsible for the formation of α-granule precursor vesicles, whereas NBEAL2 is required for maturation to α-granules. It is anticipated that MCS formation between ER and endosomes/α-granules may play important roles in mediating the functions of VPS33B/16B and NBEAL2 as ER and endosomes are repeatedly reported to be tightly associated.

Figure 1.10. Model of α-granule biogenesis. α-granules contain both endocytosed and MK- synthesized proteins. Following endocytosis, exogenous α-granule proteins are first trafficked to early endosomes located near the cell periphery. At this juncture, proteins within early endosomes undergo sorting where selective proteins can be returned to the cell surface via recycling endosomes, delivered to other cellular compartments or retained within early endosomes as it matures into late endosomal MVB I. Alternatively, α-granule cargo proteins are thought to enter α-granules directly, though the mechanisms involved are unclear. The trafficking of endogenously synthesized α-granule proteins initiates at TGN. VPS33B/16B act to facilitate the trafficking of α-granule proteins from TGN to MVB I and further promotes the maturation of MVB I into MVB II, a process exemplified by the appearance of electron-dense matrices. Mature MVB II then give rise to δ-granules, requiring the action of VPS33A/16A, and α-granules. Specifically, VPS33B/16B first assembles the precursor α-granules, marked by the presence of P-selectin, where NBEAL2 then functions in the maturation of α-granules. Figure adapted and reprinted with permission (Chen, Lo et al. 2017).

1.4 NBEAL2: a protein with unknown functions 1.4.1 BEACH domain protein family

The linking of NBEAL2 mutations to GPS increased interest in NBEAL2, for which no functions had previously been identified. NBEAL2 is a large protein, 2754 amino acids (aa, 302 kDa) long, with a complex structure containing armadillo-like (ARM) domains, a conacavalin-A like lectin (ConA) domain, pleckstrin homology (PH) domain, Beige Chediak- Higashi (BEACH) domain and WD40 domains (Gunay-Aygun, Falik-Zaccai et al. 2011, Kahr, Hinckley et al. 2011). Fig. 1.11 shows the functional homology domains of NBEAL2 and missense mutations that have being reported to cause GPS, indicating amino acid residues of critical functional importance. The BEACH domain is conserved within a family of nine human BEACH domain containing proteins (BDCP): lysosomal trafficking regulator (LYST), neurobeachin (NBEA), neurobeachin-like 1 (NBEAL1), neurobeachin-like 2 (NBEAL2), lipopolysaccharide-responsive, beige-like anchor protein (LRBA), WD and FYVE zinc finger domain containing protein 3 (WDFY3), WD and FYVE zinc finger domain containing protein 4 (WDFY4), neutral sphingomyelinase activation-associated factor (NSMAF) and WD repeat domain 81 (WDR81) (Cullinane, Schaffer et al. 2013).

The function of the BEACH domain is currently unknown. From a yeast-two-hybrid study of the BEACH domain from LYST it was postulated to function in the facilitation of protein- protein interaction to promote vesicular trafficking events (Tchernev, Mansfield et al. 2002). Results from a recent study also support the scaffolding role of the BEACH domain from NBEAL2, which binds to DOCK7 and VAC14 (Mayer, Jasztal et al. 2018). Directly preceding the BEACH domain in seven of the nine human BDCP is a PH domain, which is generally known to interact with phospholipids (Maffucci and Falasca 2001, Jogl, Shen et al. 2002). However the phospholipid interaction is not necessarily true for PH-BEACH domains (Lenoir, Kufareva et al. 2015). The PH-BEACH domains in NBEA and LRBA do not bind phospholipids, (Jogl, Shen et al. 2002, Gebauer, Li et al. 2004), while those in WDFY3 and NSMAF have been reported to be capable of binding phospholipids, suggesting functional heterogeneity among BDCP (Simonsen, Birkeland et al. 2004, Haubert, Gharib et al. 2007) . Crystallography studies of the PH-BEACH domain from NBEA and LRBA reveal that the PH domain interacts with the BEACH domain to form a single structure. It is suggested that in this conformation, PH-BEACH functions as a single unit instead of as independent domains and as such, the function of the PH domain in PH-BEACH may deviate from its classical function. Interestingly, the presence of a large groove between the PH and BEACH domain

38 suggests a potential ligand binding site (Jogl, Shen et al. 2002, Gebauer, Li et al. 2004). All members of the BDCP family contain a series of WD40 repeats following the BEACH domain. From the assembly of five to seven blades, multiple WD40 repeats form a β-propeller structure that serves as a scaffold for protein-protein interactions and protein complex assemblies (Xu and Min 2011). In six of the nine identified human BDCPs, a ConA domain, with functions in oligosaccharide binding, is present and precedes the PH-BEACH domain (Burgess, Mornon et al. 2009). ARM domains are commonly shared by BDCP for which NBEAL2 contains two ARM domain near the N-terminal, separated by the Con-A domain. Similar to WD40 repeats, ARM domains serve as a platform to facilitate protein-protein interactions (Hatzfeld 1999, Madhurantakam, Varadamsetty et al. 2012). A schematic diagram aligning the domains of the nine identified human BDCP is shown in Fig. 1.12.

BDCP proteins are not well understood and their association with disease has provided many insights, most notably that their primary functions appear to be linked to vesicular trafficking and organelle biogenesis (Cullinane, Schaffer et al. 2013). The best-studied BDCP is LYST, where mutations cause CHS, characterized by enlarged lysosomes in leukocytes and paucity of platelet δ-granules, hence LYST is postulated to be involved in lysosomal membrane trafficking via the regulation of fusion/fission events (Kaplan, De Domenico et al. 2008). NBEA is the second most studied BDCP. Mutations in NBEA are associated with autism (Castermans, Wilquet et al. 2003) and NBEA deficiency is linked to unregulated secretion of large dense core vesicles (LDCV) from neurons into synapses, resulting in impairment of neuronal function (Castermans, Volders et al. 2010). While NBEA is predominately expressed in brain tissues (Albers, Cvejic et al. 2011, Lauks, Klemmer et al. 2012), mutations also manifest in abnormal morphology of platelet δ-granules, suggesting a potential role of NBEA in platelet development (Castermans, Volders et al. 2010). Only a heterozygous mutation of NBEA has been reported clinically, indicating that NBEA is essential for survival, as indicated by mouse studies where homozygous NBEA mutations resulted in paralysis and perinatal death due to failure of synaptic transmission (Su, Balice-Gordon et al. 2004, Medrihan, Rohlmann et al. 2009).

NBEAL1 and NBEAL2, are the least studied BDCP in the family. Phylogenetic analysis indicates that NBEAL1 and NBEAL2 are products of a gene duplication event unique to vertebrates (Kahr, Hinckley et al. 2011). Sequence analysis of NBEAL1 revealed a putative vacuolar-targeting motif, which may indicate localization at lysosomes, and analysis of

NBEAL1 expression showed high levels in brain, kidney, prostate and testis (Chen, Lu et al. 2004). LRBA deficiency is frequently implicated in immune deficiency and autoimmunity (Lopez-Herrera, Tampella et al. 2012). Situated at the RAB11-positive recycling endosomes in T-cells, LRBA acts to prevent the lysosomal targeting of internalized cytotoxic T lymphocyte antigen–4 (CTLA-4), a surface receptor that prevents autoimmune responses. This prevents CTLA-4 degradation and allows it to be recycled back to the cell surface (Lo, Zhang et al. 2015). These observations suggest that LRBA functions as a scaffold protein in vesicular trafficking events (Sansom 2015). Ubiquitously expressed, WDFY3 is localized to autophagosomes (Simonsen, Birkeland et al. 2004). In contrast to the PH-BEACH domain from NBEA and LRBA which is reported to be incapable of binding phospholipids (Jogl, Shen et al. 2002, Gebauer, Li et al. 2004), WDFY3 can bind to phosphatidylinositol 3-phosphate (PI3P) (Simonsen, Birkeland et al. 2004). WDFY3 is involved in the selective autophagy of protein aggregates and functions as a scaffold protein in this process (Yamamoto and Simonsen 2011). Different from the ubiquitous expression of WDFY3, WDFY4 is expressed primarily in immune tissues and is postulated to be implicated in autoimmunity (Yang, Shen et al. 2010).

NSMAF is one of the two BDCP that lacks both the PH and ConA domain. The PH domain in NSMAF is replaced by a GRAM domain, also known to function in phospholipid binding (Doerks, Strauss et al. 2000, Tsujita, Itoh et al. 2004). NSMAF participates in immunity by mediating the tumor necrosis factor (TNF) pathway via interaction with the TNF receptor (Adam-Klages, Adam et al. 1996). This interaction promotes the signaling of pro-apoptotic events by activating downstream caspase 8 (Segui, Cuvillier et al. 2001). It is also suggested that NSMAF can mediate TNF-induced inflammation (Mas, Danjoux et al. 2013). Similar to WDFY3, NSMAF can bind to phospholipids. Through its GRAM domain, initially thought to be PH domain, NSMAF binds to phosphatidylinositol 4,5-bisphosphate (PI4,5P) at the plasma membrane, a result that is consistent with its interaction with the TNF receptors near the cell surface (Haubert, Gharib et al. 2007). WDR81 is one of the two BDCP that lacks both the PH and ConA domains and its gene mutation is implicated in cerebellar ataxia, mental retardation, and dysequilibrium syndrome (CAMRQ) (Gulsuner, Tekinay et al. 2011).

Figure 1.11. Diagram of NBEAL2 displaying homology domains and GPS-causing mutations. Schematic diagram of the functional domains of NBEAL2. Marked are the missense mutations that have being reported to cause GPS, revealing functionally important amino acids. Nonsense mutations are not shown.

Figure 1.12. Schematic of the domains of BDCP. Diagram aligning the known domains of the nine identified human BDCP. Note that BDCP is a family of large proteins, each harboring multiple functional domains. These features provide reasonable basis in support of the BDCP that were postulated to function as protein scaffolds. Diagram to scale and bar = 250 amino acid residues. Figure adapted and reprinted with permission (Cullinane, Schaffer et al. 2013).

1.4.2 NBEAL2 in megakaryocytes

While the function of NBEAL2 was unknown prior to its association with GPS, recent studies have begun to shed some light. NBEAL2 is predominately expressed in hematopoietic cells and is highly upregulated during megakaryopoiesis (Albers, Cvejic et al. 2011). Recently, it has been shown that the expression of NBEAL2 is regulated by the transcription factor, GATA1, through an enhancer interaction upstream of NBEAL2 (Wijgaerts, Wittevrongel et al. 2017). As mentioned previously , GATA1-deficiency has been linked to impairment of megakaryopoiesis and thrombopoiesis (Shivdasani, Fujiwara et al. 1997, Tsang, Visvader et al. 1997, Takahashi, Komeno et al. 1998, Mehaffey, Newton et al. 2001, Chang, Cantor et al.

2002, Crispino 2005). Since NBEAL2-deficient MKs also display developmental defects, one compelling hypothesis is that GATA1 may regulate megakaryopoiesis through the action of NBEAL2. GPS is characterized by bleeding diathesis and thrombocytopenia. This is consistent with the silencing of NBEAL2 in zebrafish, which caused the loss of thrombocyte production leading to spontaneous bleeding (Albers, Cvejic et al. 2011), although the connection is tenuous since only mammals generate megakaryocytes.

To investigate the role of NBEAL2 in megakaryopoiesis and α-granule biogenesis, NBEAL2 knockout mouse strains have been developed. The one used in this thesis recapitulates the GPS phenotype in having absent platelet α-granules and greatly decreased granule contents (Deppermann, Cherpokova et al. 2013, Kahr, Lo et al. 2013), as well as prolonged bleeding, impaired platelet function and thrombocytopenia (Deppermann, Cherpokova et al. 2013, Kahr, Lo et al. 2013, Guerrero, Bennett et al. 2014). Our studies with this Nbeal2-/- strain have revealed that NBEAL2 is required for MK maturation (Kahr, Lo et al. 2013), a result supported by other studies (Guerrero, Bennett et al. 2014) that also showed the lifespan of Nbeal2-/- platelets is normal (Deppermann, Cherpokova et al. 2013). Nbeal2-/- MK display decreased proplatelet production (Kahr, Lo et al. 2013), and it is postulated that delayed MK maturation and decreased proplatelet production are fundamental causes of thrombocytopenia in Nbeal2-/- mice. We also identified a role for NBEAL2 in α-granule biogenesis by examining the fate of cargo proteins in Nbeal2-/- MKs (Kahr, Lo et al. 2013).

Recently, it was reported that NBEAL2 interacts with DOCK7 (a guanidine exchange factor), VAC14 (a PI3,5P regulator) and SEC16 (an ER resident protein). DOCK7 expression is significantly reduced in Nbeal2-/- platelets, and DOCK7 shows reduced binding to NBEAL2 containing GPS-causing mutations, and it has been proposed that NBEAL2-DOCK7 interactions may be important in platelet formation and function (Mayer, Jasztal et al. 2018). While the mechanisms by which NBEAL2 participates in α-granule biogenesis are still unclear, the potential binding of NBEAL2 to SEC16, an ER membrane protein, and VAC14, a phospholipid regulator, raises some interesting possibilities.

1.5 Rationale, Hypothesis and Aim

Platelets play important roles in numerous physiological processes and α-granules are of major importance to platelet function, yet little is known about the mechanisms of α-granule biogenesis and regulation. NBEAL2 has recently been linked to α-granule biogenesis. Specifically, NBEAL2 deficiency results in the loss of α-granule cargo with the retention of granular membrane structures containing P-selectin. Several BDCPs have been identified as scaffold proteins involved in vesicular trafficking events, and we hypothesize that NBEAL2 acts as a scaffold during the trafficking and targeting of cargo proteins to nascent α-granules, facilitating their maturation and subsequent secretory function. The aim of this study was to investigate the role of NBEAL2 in α-granule biogenesis, and several approaches were employed. The molecular and structural properties of NBEAL2 were examined via biochemical analysis. Putative NBEAL2 interaction partners were identified to provide insights into cellular processes this protein may be involved with. This approach will place emphasis on interactions that may be linked to α-granule formation and function. The consequences of NBEAL2 loss were examined in knockout mice and platelets and MKs derived from them, with comparisons to normal animals and cells revealing potential roles for NBEAL2 in α- granule biogenesis and function, and in MK development and platelet production.

Chapter 2

Materials and Methods

2.1 Human megakaryocytes and platelets

Primary human megakaryocytes were generally derived from CD34+ hematopoetic progenitors, refer to Section 2.3.1 for details. In instances where hematopoetic progenitors were obtained from human bone marrow aspirates or peripheral blood, density gradient centrifugation was first performed to obtain the mononuclear cell populations which contain the hematopoetic progenitors. Samples were collected into 3.2% citrate to prevent coagulation. Samples were layered onto Ficoll-Paque (GE Healthcare) in a volume ratio of 1:1 and centrifuged (400g, 30 minutes) without brake. Following centrifugation, five distinct layered fractions became apparent. The uppermost layer was aspirated and the second fraction containing the mononuclear cells were transferred to a new vessel. The cells were washed 3 times with PBS/ACD (pH 6.1) with centrifugation at 200g for 10 minutes per wash. The cells were resuspended and cultured in IMDM (Gibco) supplemented with 10% BIT 9500 (Stemcell Technologies), 2 mM l-glutamine (Wisent), 1% penicillin-streptomycin (Wisent) and 50 ng/mL thrombopoietin (a gift from Kirin Brewery Company), 50 ng/mL of stem cell factor

(at 37°C and 5% CO2 for 12 days for differentiation into megakaryocytes. Human platelets were obtained through isolation from human whole blood, refer to Section 2.3.4 for details.

2.2 Knockout and transgenic mice

Nbeal2-/- mice were produced from spermatozoa of the strain B6;129S5-Nbeal2tm1Lex/Mmucd obtained from the Mutant Mouse Regional Resource Center at the University of California, Davis. Knockout was achieved by deleting exons 4 to 11 of the 54-exon NBEAL2 gene through homologous recombination. The spermatozoa were used to fertilize C57BL/6J oocytes where the embryo was then transferred into a WT C57BL/6J female to generate progenies. Homozygous Nbeal2-/- mice were obtained through cross-breedings. The knockout was confirmed by analyzing the deletion site via PCR, specifically, with 1 5’ primer (5’- GTCCTGCTTGACCTACCGTC-3’) and 2 3’ primers (5’- CAGGGAGGATAACGAGATAGTCTT-3’ and 5’-CCTAG GAATGCTCGTCAAGA-3’). Two PCR reactions using the 5’ primer with one of the 3’ primer yielded the predicted products: 223 bp and no product for WT and no product and 401 bp for Nbeal2-/-.

To rescue the GPS phenotypes observed in Nbeal2-/- mice and to better understand the dynamics of NBEAL2 expression, NBEAL2 was transgenically introduced into Nbeal2-/- mice with its expression being regulated by a megakaryocyte-specific promoter, aIIb. To generate

45 the transgene rDNA, PLJM1-3XFlag-EGFP-NBEAL2 was first generated from modified PLJM1-EGFP vector (Addgene plasmid #19319). PLJM1-EGFP was modified as the EGFP sequence was deleted using AgeI-EcoRI restriction digests and replaced with sequences containing AgeI-3XFlag-SalI-MluI-PacI. EGFP was subcloned into the construct via SalI-MluI restriction sites. NBEAL2 was then subcloned into the vector via MluI-PacI restriction sites to generate PLJM1-3XFlag-EGFP-NBEAL2. CMV promoter from the resulting construct was excised using NdeI-NheI restriction digests and replaced with aIIb promoter (a gift from Dr. David A. Wilcox from Children's Hospital of Wisconsin Research Institute, Milwaukee, WI, USA). SV40 PolyA was added to the C-terminal of 3XFlag-EGFP-NBEAL2 using PacI-BamHI restriction digests. The transgene rDNA was prepared by digesting the resulting plasmid constructs using NdeI and BamHI. Starting from the 5’ end, the rDNA contains sequences for allb promoter, 3XFlag, EGFP, NBEAL2 then SV40 PolyA. The rDNA was microinjected into Nbeal2-/- embryos and the progenies were then examined for the presence of transgene.

2.3 Cell lines and cultures

Cell lines and cultures used are compiled in the following table.

Cell Line Type Source Product DAMI Human Megakaryocytes Gift from Dr. D. Wilcox N/A ImMKCL Human Megakaryocytes Gift from Dr. Koji Eto N/A HEK293 Human Embryonic ATCC CRL-1573 Kidney HEK293T Human Embryonic ATCC CRL-3216 Kidney HEK293F Human Embryonic Thermo Fisher R79007 Kidney AH109 Yeast-Two-Hybrid Clontech 630489 Y187 Yeast-Two-Hybrid Clontech 630489 DH5α Bacterial Cloning Thermo Fisher 18265017 STBL3 Bacterial Cloning Thermo Fisher C737303 BL21(DE3) Bacterial Protein Millipore 69450 Expression BL21(DE3) Star Bacterial Protein Thermo Fisher C601003 Expression Rossetta (DE3) Bacterial Protein Millipore 70954 Expression

2.3.1 Isolation and culture of human primary CD34+ cells

Cultured primary human megakaryocytes were differentiated from CD34+ hematopetic progenitor cells. CD34+ cells were obtained from G-CSF–mobilized blood from bone marrow

46 transplant donors through affinity purification using the CD34 MicroBeads kit (Miltenyi Biotec). To prevent column overloading, mononuclear cells were enriched with Ficoll density gradient prior to CD34 affinity purification as described in section 2.1 when sample size was greater than 1mL. CD34+ cells were cultured with IMDM (Gibco) supplemented with 10% BIT 9500 (Stemcell Technologies), 2 mM l-glutamine (Wisent), 1% penicillin-streptomycin (Wisent) and

50 ng/mL thrombopoietin (a gift from the Kirin Brewery Company) at 37°C and 5% CO2 for up to 12 days to allow time for differentiation and maturation. For immunofluorescence, MKs from day 8 of culture were seeded onto matrigel-coated coverslips (BD Biosciences, 1:6~1:10 dilutions) in 12-well plates and fixed on day 12.

2.3.2 Isolation and culture of mouse primary bone marrow cells

Primary bone marrow cells were flushed from mice femurs and tibias and filtered through 100µm mesh nylons to remove the debris. Cells were washed 2 times with PBS via centrifugation (200g, 10 min) and resuspended in DMEM (Wisent Inc.) supplemented with 10% FBS (GIBCO),1% Penicillin-Streptomycin (Wisent Inc.) and 50 ng/mL of rhTPO (gift from Kirin Brewery Company). Alternatively, cells were resuspended in IMDM (Gibco) supplemented with 10% BIT 9500 (Stemcell Technologies), 2 mM l-glutamine (Wisent), 1% penicillin-streptomycin (Wisent) and 50 ng/mL rhTPO (a gift from the Kirin Brewery Company).

Cells were incubated (37°C, 5% CO2) for indicated days then seeded onto matrigel-coated coverslips (BD Biosciences, 1:6~1:10 dilutions) in 12-well plates.

Seeded cells were fixed at indicated times and prepared for immunostaining. MK population expansion was determined by comparing cytospun cells from native bone marrow and day 5 culture. MKs were visualized through CD41 stains following fixation. Cells were imaged at 4x and total numbers were enumerated by counting DAPI-stained nuclei; MKs were scored as CD41-positive cells. Relative ratios of CD41 positive and DAPI positive cells were calculated using Image Pro 6 (Media Cybernetics) software.

2.3.3 Enrichment and purification of human and mouse primary megakaryocytes

Enrichment of MKs was performed using BSA gradient. Cultured primary human and mouse MKs were washed 3 times with PBS via centrifugation (200g, 10 min) and resuspended in 3 mL of PBS. BSA gradient was prepared by carefully layering 3mL of 1.5% BSA in PBS on top of 3 mL of 3% BSA in PBS in a 15 mL tube. 3 mL of cells in PBS were carefully layered on top of the BSA gradient and incubated (37°C, 5% CO2) for 30min to allow for cell

47 sedimentation. Supernatants were removed and the enriched cell pellet was washed with PBS and resuspended into appropriate media. Alternatively, mouse MKs were affinity purified using mouse and rat CD61 MicroBeads kit (Miltenyi Biotec) according to the manufacturer’s instructions and resuspended into appropriate media for downstream applications.

2.3.4 Isolation of human and mouse platelets

Whole blood from human donors or mice were collected into 3.2% citrate and centrifuged at room temperature without brake (100g, 10 min). The upper layer was carefully removed and collected as the platelet-rich plasma (PRP). Platelets were pelleted via centrifugation at room temperature (1000g, 10 min) and gently washed 3 times with PBS/ACD (acid citrate dextrose, pH 6.1). The washed platelets were resuspended in PBS at a concentration of at a concentration of 1*109 platelets/mL. Cells were lysed for 10 min at room temperature with the addition of Triton X-100 (0.5% final) and protease inhibitor cocktail (Roche, 2X final). Debris was removed via centrifugation (16,000g, 10 min) and the supernatants were stored as platelet lysates.

2.4 Antibodies

Summary of primary antibodies used for immunofluorescence (IF), immunogold (IG) and immunoblot analysis (IB) with concentrations.

Type Antigen Dilution IF Dilution IG Dilution IB Source Product

RABBIT EEA1 200 Cell Signaling C45B10 CD71/TFR 200 Abcam ab84036 ETV6 100 Thermo Scientific PA5-35371 FGN 20 Dako A0080 FLAG 100 1000 Sigma F7425 HA 1000 Bethyl A190-208A NBEAL2 100 10 1000 Abcam ab187162 RAB4 100 Thermo Fisher PA3-912 RAB5 200 Cell Signaling C8B1 RAB7 100 Cell Signaling D95F2 RAB11 100 Cell Signaling D4F5 RAB27B 100 Millipore ABS1026 SEC22B 100 1000 Synaptic Systems 186003 TSP1 200 2000 RayBiotech MD-01-0027 VAMP2 1000 Cell Signaling D601A VAMP8 200 Synaptic Systems 104302

VWF 400 5 Dako A0082

MOUSE AP-1 50 Sigma A4200 CALNEXIN 1000 Abcam ab31290 CD61 200 Dako M0753 CD63 100 Hybridoma Bank H5C6 CD71/TFR 100 Sigma C2063 CLATHRIN 400 BD Biosciences 610500 FLAG 100 1000 Sigma F1804 GAPDH 5000 Millipore MAB374 GM130 50 BD Biosciences 610823 HIS 1000 Medimabs MM-0165-P LAMP1 200 Abcam H4A3 MYC Covance MMS-150R RAB4 100 BD Biosciences R68520 RAB5 50 BD Biosciences 610725 RAB11 100 BD Biosciences 610657 TSP1 500 Thermo Fisher MA5-13385 TSP1 100 R&D Systems MAB3074 TRANSFERRIN 1000 GeneTex N3C3 α-TUBULIN 5000 Sigma-Aldrich T6074 β-ACTIN 5000 Sigma-Aldrich A5441

GOAT P-SELECTIN 100 1000 Santa Cruz sc-6941

SHEEP FGN 200 1000 Affinity Biologicals SAFG-AP RAB27A 100 R&D Systems AF7245 TGN46 500 BioRad AHP500 VWF 2000 1000 ABD Serotech AHP062

RAT LAMP1 1000 Abcam 1D4B CD41 200 eBioscience 14-0411

2.5 Constructs 2.5.1 Bacterial expression constructs

Subclonings were performed using DH5α bacteria (Thermo Fisher).

PH-BEACH domain (1918-2345aa) of NBEAL2 was subcloned into pET28a vector (Millipore) via BamHI-XhoI restriction sites, pETDuet-1 vector (Millipore) via BamHI-NotI restrictions sites.

The same PH-BEACH domain was subcloned into pET His6 TEV LIC cloning vector (1B, addgene #29653), pET His6 NusA TEV LIC cloning vector (2N-T, addgene #29709), pET His6 Sumo TEV LIC cloning vector (2S-T, addgene #29711) and pET His6 MBP TEV LIC cloning vector (1M, addgene #29656) using ligation independent cloning (LIC) according to the instructions.

2.5.2 Yeast expression constructs

Subclonings were performed using DH5α bacteria (Thermo Fisher).

N-terminal (1-1470aa) and C-terminal (1404-2724aa) of NBEAL2 were subcloned into pGBKT7 via NdeI-BamHI restriction.

2.5.3 Mammalian expression constructs

Subclonings were performed using DH5α bacteria (Thermo Fisher).

To utilize the piggyback system (Li, Michael et al. 2013), PH-BEACH domain (1918-2345aa) of NBEAL2 was subcloned into PB-T-PAF (a gift from Dr. James Rini, Department of Biochemistry, University of Toronto) via AscI-NotI restriction sites using DH5α.

NBEAL2 was subcloned into p3XFlag-CMV-14 using NotI-BamHI restriction digests. EGFP was then added to the N-terminal of NBEAL2. The expressed NBEAL2 harbors EGFP at the N-terminal and 3XFlag at the C-terminal.

To generate HA-NBEAL2, full-length NBEAL was subcloned into pCMV-HA vector (Clontech) using the XhoI-NotI restriction sites.

SEC22B cDNA was obtained from Origene and subsequently subcloned into p3xFLAG-CMV- 14 vector (Sigma) via NotI-BamHI restriction sites using DH5α.

Fragments of NBEAL2 were subcloned into pCMV-HA vector (Clontech) using XhoI-NotI restriction sites: C1 (1-550aa), C2 (530-903aa), C3 (877-1403), C4 (1380-1903aa), C5 (1877- 2400aa), C6 (2384-2754aa), C7 (1-1145aa), C8 (1798-2754aa), C9 (1380-2754aa), C10 (1380-1797aa), C11 (1798-1903aa). C10 was also subcloned into pCMV-myc vector (Clonetech) using Xho-NotI restriction sites.

2.5.4 Lentiviral constructs

Subclonings involing lentiviral constructs were performed in RecA- STBL3 bacteria (Thermo Fisher) to ensure the stability of these constructs.

PLJM1-3XFlag-EGFP-NBEAL2 was generated from modified PLJM1-EGFP vector (Addgene plasmid #19319). PLJM1-EGFP was modified as the EGFP sequence was deleted using AgeI-EcoRI restriction digests and replaced with sequences containing AgeI-3XFlag-SalI- MluI-PacI. EGFP was subcloned into the construct via SalI-MluI restriction sites. NBEAL2 was then subcloned into the vector via MluI-PacI restriction sites to generate the final construct.

To generate CRISPR knockout cell lines via lentiviral transduction, guide RNA (gRNA) oligos were cloned into lentiCRISPR v2 (addgene #52961). LentiCRISPR plasmid was prepared with gel-extraction following digestion with BsmBI (New England Biolabs) and treatment with alkaline phosphatases (New England Biolabs). Guide RNA were designed using MIT CRISPR Design (Zhang lab). Guide oligos containing BsmBI-compatible ends were phosphorylated with T4 PNK (New England Biolabs) and ligated onto BsmBI-digested lentiCRISPR plasmid with T4 DNA ligase (New England Biolabs). The ligated products were then transformed into STBL3 bacteria.

To generate GPS mutations and rescue constructs, NBEAL2 E1833K or R1839C mutations were generated via site-specific mutagenesis from pCMV-HA-NBEAL2. Full-length NBEAL2, NBEAL2-E1833K and NBEAL2-R1839C were subsequently subcloned into pLenti CMV Blast empty (w263-1, addgene #17486) via SalI-BamHI restriction sites. A HA-tag was added to the N-terminal end NBEAL2 during this process. SEC22B was also subcloned into the same vector via SalI-BamHI restriction sites where a HA-tag was added to its N-terminal end.

2.6 Transfection and electroporation

For general purpose transfections, Lipofectamine 2000 (Thermo Fisher) and jetPRIME (Polyplus transfection) were used. For transient transfection, 1x106- 2x106 HEK293/HEK293T cells seeded onto a 10 cm dish in 10 mL culture media (DMEM supplemented with 5% FBS) the day before transfection to achieve a confluency of 60-80% on the day of transfection. 5- 10 μg of DNA was mixed with 500 μL of jetPRIME buffer and 20 μL of jetPRIME reagents, incubated at room temperature for 10 min then added to the 10 cm plate. The cells were

51 harvested 48-72 h post-transfection for analysis. For DAMI cells, 1x106 cells were seeded onto 6-well plate in 2 mL of culture media (IMDM supplemented with 5% horse serum) prior to transfection. 10 μL of Lipofectamine 2000 was added to 250 μL of OptiMEM (Thermo Fisher) in a tube and 4 μg of plasmid DNA was added to 250 μL of OptiMEM in another tube. The tubes were incubated for 5min, combined and incubated for another 20 min before addition to the seeded DAMI culture. Gene expression was analyzed 48-72 h post-transfection.

For transfection of human primary CD34+ cells or to achieve a higher plasmid transfer efficiency in DAMI cells, Nucleofector Kits for Human CD34+ Cells (Lonza) was used. 5x106 cells were resuspended in 100 μL of Nucleofector solution with 5 μg of plasmid DNA and transferred into an electroporation cuvette. Cuvette was inserted into a Nuclecfector system (Lonza) and program U-008 was used to enable the electroporation. Cells were immediately and gently transferred into 2 mL of prewarmed media for recovery. Cells were assayed for gene expression 48-72 h post-electroporation.

2.7 Lentivirus production and transduction 2.7.1 Protein expression in mammalian cell lines and human primary CD34+ cells

1x106- 2x106 HEK293T cells were seeded onto a 10 cm dish in 10 mL culture media (DMEM supplemented with 5% FBS) the day before transfection to achieve a confluency of 60-80% on the day of transfection. To prepare the transfection mixture, 3 mL of OptiMEM (Thermo Fisher) was mixed with 20 ug of expression vector, 12 μg of pMD2.G (addgene #12259), 20 μg pMDLg/pRRE (addgene #12251), 10 μg pRSV-Rev (addgene #12253) and 70μL of Lipofectamine 2000 (Thermo Fisher) and incubated for 20 min at room temperature before addition to the HEK293T cell culture. Media was exchanged for fresh culture media containing 1mM of sodium butyrate 12 h post-transfection. Supernatants were collected every 24 h post-transfection for 72 h. The supernatants were pooled, centrifuged (500g, 5min) to remove debris and filtered through a 0.45 μm mesh. Viruses were pelleted via ultracentrifugation (100,000g, 2 h, 4°C), resuspended in PBS and stored at -80°C.

CD34+ cells, DAMI cells and imMKCL cells were transduced with lentivirus using the following protocol. CD34+ cells were isolated as described in section 2.3.1.

To perform virus transduction, cells were seeded onto 24-well plates at 70% confluency. Culture media were aspirated and 1 mL of culture media containing 8ug/mL of polybrene and

52 viruses were added to the wells. Transduction efficiency was assayed 48 h post-transduction via western blots or GFP detection.

Alternatively, cells were transduced with a higher efficiency via retronectin. 24-well non-tissue culture treated plates (Falcon) were coated with 300 μL of 0.1 mg/mL of retronectin (Clontech) per well for 2 h at room temperature or overnight at 4°C. The wells were then blocked with 300 μL of 2%BSA in PBS for 30min at room temperature and washed with 600μL of Hank’s balanced salt (Gibco) supplemented with 2.5% (v/v) 1 M Hepes. 1.5*106 cells were resuspended in 500 uL of culture media, mixed with 20 μL of viruses, added to the coated wells and incubated (37°C, 5% CO2). On the next day, the media was replaced with fresh media containing 20 μL of viruses and incubated for another day. Media was removed and the cells were collected from the wells through forceful PBS washes, trypsin or cell dissociation buffer (GIBCO). Cells were washed 3 times with PBS (200g, 5 min) and resuspended in fresh culture media.

2.7.2 CRISPR knockout in mammalian cells

To generate CRISPR knockout, LentiCRISPR v2 cloned with a gene-specific gRNA was used as the expression vector to generate the virus as described in section 2.7.1. Specifically, protein expressions in imMKCL cells were knocked-out through CRISPR. ImMKCL cells, a megakaryocytic cell line, were a gift from Dr. Koji Eto from Japan (Nakamura, Takayama et al. 2014). ImMKCL cells were grown in hematopoetic differentiation medium for maintenance (IMDM (Gibco) containing 10 μL/mL of Insulin-Transferrin-Selenium 100X (Thermo Fisher), 2 mM of L-glutamine (Thermo Fisher), 0.45 mM of α-monothioglycerl (Sigma), 50 ug/mL of ascorbic acid and (Sigma), 10% FBS, 50 ng/mL of thrombopoietin (a gift from Kirin Brewery Company), 50 ng/mL of human stem cell factor (Millipore) and 1 ug/mL of doxycycline (Wisent). ImMKCL cells were transduced with retronectin-coated plates as described in section 2.7.1. Cells were selected for successful transduction via puromycin selection at 1ug/mL. Surviving colonies were then expanded. Prior to assay, cells were wash 3 times with PBS (400g, 5min) and resuspended in hematopoetic differentiation medium without doxycycline to initiate differentiation. The cell medium was changed again the next day, incubated (37°C, 5% CO2) for 5 days, then assayed for knockout with western blot.

2.8 Yeast two-hybrid

N-terminal and C-terminal NBEAL2 pGBKT7 constructed were generated as described in section 2.5.2. The constructs were transformed into AH109 yeast cells as baits. AH109 cells were grown in YPDA media (yeast extract, peptone, dextrose, adenine containing 20 g/L peptone, 10 g/L yeast extract, 20 g/L glucose, 0.1 g/L NaOH and 0.003% adenine hemisulfate) at 30°C to an OD600 of 0.5-0.7, pelleted (1500g, 5min), resuspended in 1.5mL of TE/LiOAc buffer (10 mM Tris, 1 mM EDTA, 0.1 M LiAc, pH 7.5) and divided into 100 μL transformation aliquots. 50 μg of salmon sperm DNA (Thermo Fisher) was boiled at 95°C for 5min and mixed with 100 ug of bait construct DNA before being added to a 100μL transformation aliquot. 300 μL of TE/LiOAc/PEG-3350 buffer (10 mM Tris, 1mM EDTA, 0.1M LiAc, 50% w/v PEG-3350, pH7.5) was subsequently added to the transformation aliquot and incubated at 4°C for 30min. 70 μL of DMSO was added to the reaction followed by incubation at 42°C for 15 min. To select for transformed colonies, the reaction mixture was spread on a selection plate that contains synthetic dextrose medium but lacks tryptophan (SD/-Trp containing 6.7 g/L of yeast nitrogen base without amino acids, 0.6 g/L of -Trp dropout mix, 20 g/L of glucose, 20 g/L of granulated agar) and incubated at 30°C for 3 days. Colonies were then assayed for bait protein expression and auto-activation. Adequate AH109 colonies were grown to an OD600 of 0.8 in SD/-Trp media, pelleted (1500g, 5 min), and resuspended in 50 mL of 2X YPDA media. A vial of human bone marrow library (Clontech), containing Y187 yeast cells that harbor an assortment of prey proteins resembling the bone marrow proteomics, were thawed at 37°C and added to the same 2X YPDA media culture. The yeast culture was then incubated for 16h with shaking (50rpm). Following mating, cells were pelleted (1500g, 10min) and resuspended in 10mL of 0.5X YPDA. In 250μL aliquots, the mated yeasts were spread onto quadruple amino acid dropout plates (QDO containing SD/- Ade/-His/-Leu/-Trp and 20mg/mL of x-α-gal (Clontech)) and incubated at 30°C for 1 to 2 weeks. To ensure stringency, positive blue colonies were picked and grown on a fresh QDO plate for 5 days. Prey DNA sequences from each positive colony were identified using Matchmaker Insert Check PCR Mix 2 (Clontech) according to the manufacturer’s instructions. Alternatively, plasmids from the positive colonies were isolated through phenol/chloroform/isoamyl extraction. Briefly, a 5 mL yeast culture pellet was resuspended in 150 μL of plasmid isolation buffer (2% Triton X-100, 1% SDS, 100 mM NaCl, 10 mM Tris and 1mM EDTA). 400 μL of phenol/chloroform/isoamyl (Sigma) and 0.3 g of glass beads were subsequently added. The mixture was vortexed for 2 min, centrifuged (16,000g, 5 min), and the upper aqueous layer containing the plasmid DNA was collected. For further extraction, an

54 addition 150 μL of plasmid isolation buffer was added, mixed, centrifuged (16,000g, 5 min) and the upper aqueous layer was transferred and combined with the previous fraction. Prey sequences were amplified with PCR and sequenced. The prey proteins were identified using matching DNA sequences through Basic Local Alignment Search Tool (BLAST).

2.9 Western blotting

Cell lysates were prepared by incubating cell pellets with cold lysis buffer (50 mM Tris, 137 mM NaCl, 1 mM EDTA, 1% Triton X-100, pH 8, 1X protease inhibitor cocktail (Roche)) for 10min at 4°C. Crude lysates were centrifuged (16000g, 10min) and supernantants were stored at -20°C or added to 2X SDS sample buffer (0.13M Tris, 20% w/v glycerol, 4% w/v SDS, 0.001% w/v bromophenol blue, pH 6.8). Reduced samples were treated with 2% v/v of β-mercaptoethanol and boiled for 5 min at 95°C.

Unreduced and reduced samples were run on SDS-PAGE gels and transferred nitrocellulose membrane according to standard procedures. Blots were probed with primary antibodies and corresponding secondary antibodies. Blots were exposed with enhanced chemiluminescence substrates (ECL, Thermo Fisher) and imaged with Odyssey FC (LI-COR Biosciences). Media and lysates from MKs incubated with bio-FGN were probed with HRP-conjugated streptavidin (Thermo Fisher, 434323, dilution 1:5000) or sheep anti-FGN. Mouse anti-GAPDH and anti- Transferrin antibodies were used as sample loading controls.

Image preparation and densitometry analysis were performed using Image Studio Lite (LI- COR Biosciences).

2.10 Immunoprecipitation

HEK293 cells were transfected with jetPrime (Polyplus tranfection) as described in section 2.6. Plates were gently wash 2 times with PBS to remove residual culture media. 1 mL of cold lysis buffer (50 mM Tris, 137 mM NaCl, 1 mM EDTA, 1% Triton X-100, pH 8, 1X protease inhibitor cocktail (Roche)) was then added to each 10 cm plate, cells were scraped, collected into a tube and incubated on ice for 30 min with occasional mixing. For endogenous protein immunoprecipitation, platelets or primary megakaryocytes were isolated as previously described and resuspended in the same cold lysis buffer before incubation on ice for 30 min. Lysates were centrifuged (16000g, 10 min), 25 μL of the supernatants were added to 25 μL of

SDS sample buffer (0.13 M Tris, 20% v/v glycerol, 4% w/v SDS, 0.001% w/v bromophenol blue, 2% v/v β-mercaptoethanol, pH 6.8) as the input loading control. The remaining supernatants were collected and divided into two equal portions where 1 μg of capturing antibody was added to one portion and a negative control IgG, from the same host species as the capturing antibody, was added to the other. The reactions were incubated on a rotator for 1-2h at 4°C. 70 μL of washed agarose bead slurry (50%, Protein A (Sigma) or Protein G (GE Healthcare)) were added to the both reactions according to the host species of the capturing antibody used and incubated on a rotator for 1 h 4°C. Beads were centrifuged (10000g, 5seconds) and washed 4 times with cold wash buffer (50 mM Tris, 137 mM NaCl, 1 mM EDTA, 0.1% Triton X-100, pH 8) or cold PBS. 50 μL of 2X SDS sample buffers were added to each tube and the samples were boiled for 5min at 95°C to elute the immunoprecipitants. Samples were then loaded unto SDS-PAGE and analyzed with western blot.

2.11 Blue Native gel electrophoresis

Cell lysates were diluted with 4X NativePAGE Sample Buffer or dialyzed overnight against 1X NativePAGE Sample Buffer (50mM Bis-Tris, 50mM NaCL, 10% v/v glycerol, 0.001% Ponceau S, pH 7.2). Protein ladder, NativeMark Unstained Protein Standard (Thermo Fisher), and samples were loaded onto 4-16% NativePAGE Bis-Tris gels (Thermo Fisher), followed by electrophoresis at 150V for 3 h with NativePAGE Light Blue Cathode Buffer (50 mM Bis- Tris, 50 mM Tricine, 0.002% G-250, pH 6.8). For samples containing detergents, electrophoresis was initially performed at 150V with NativePAGE Dark Blue Cathode Buffer (50 mM Bis-Tris, 50 mM Tricine, 0.02% G-250, pH 6.8) for 1 h before the buffer was exchanged for NativePAGE Light Blue Cathode Buffer for another 2 h. Gel was transferred onto PVDF membrane with Native Transfer Buffer (25 mM Bicine, 25 mM Bis-Tris, 1 mM EDTA, pH 7.2) at 100V for 1 h or 30V overnight.

2.12 Cell fractionation

DAMI cells or primary megakaryocytes were washed 3 times with PBS (200g, 5 min), washed twice with cold homogenization buffer (25 mM HEPES, 0.25M sucrose, 1 mM EDTA, 1 mM DTT, pH 7.4, 1X protease inhibitor cocktail (Roche)), then resuspended into 300μL of cold homogenization buffer. Cells were lysed mechanically by aggressively expelling the cells through a 25-gauge syringe needle against the bottom of a 15 mL conical tube repeatedly. The lysis was repeated until cell lysis were deemed satisfactory when the lysates were examined under a light microscope (e.g. free of intact cells, approximately 20 repeats).

Lysates were centrifuged (3000g, 15 min, 4°C) to remove intact cells and debris. Supernatants were then ultracentrifuged (120,000g, 2 h, 4°C) using a TLS55 rotor. Supernatants were collected as the cytosolic fraction. Pellet was resuspended in homogenization buffer supplemented with 1% Triton X-100, incubated on ice for 15min for solubilization, and collected as the membrane fraction. The fractions were dialyzed and loaded onto Blue Native gels as described in section 2.11.

2.13 Mass spectrometry and analysis

Stable DAMI cells expressing 3XFlag-EGFP-NBEAL2 was generated by transducing DAMI cells with lentivirus packaged with PLJM1-3XFlag-EGFP-NBEAL2 expression vector. Techniques are described in section 2.5.4 and 2.7.1. 1μg/mL of puromycin was supplemented to the DAMI culture 2 days post-transduction to select for positive colonies. Surviving colonies were isolated, expanded, and assayed for the expression of 3XFlag- EGFP-NBEAL2 via western blot following stimulation with 50 ng/mL of Phorbol 12-myristate 13-acetate (PMA) for 2 days. To perform immunoprecipitation mass spectrometry, stable DAMI cells expressing 3XFlag-EGFP-NBEAL2 were grown in 15 cm dishes and stimulated with 50ng/mL of PMA for 2 days. 1x108 cells were collected via trypsinization, washed 3 times with PBS (300g, 5min), resuspended in cold lysis buffer (50 mM Tris, 137 mM NaCl, 1 mM EDTA, 1% Triton X-100, pH 8, 1X protease inhibitor cocktail (Roche)) and incubated, with occasional inversions, at 4°C for 30 min. Lysates were then centrifuged (16000g, 10 min) and the supernatants were divided into two equal portions in two vials. 150 μL of washed anti- FLAG M2 affinity gel slurry (50%, Sigma) was added to one portion and 150 μL of Cross- linked Sepharose CL-4B bead slurry (50%, Sigma) was added to the other portion as a negative control. The vials were incubated on a rotator for 2 h at 4°C. The beads were then washed 4 times (10000g, 5seconds) with cold PBS and the immunoprecipitants were eluted with 3XFlag peptide competition with 250 μg/mL of 3XFlag peptides or with 9 M NH4OH (pH 11-12) supplemented with 0.5 mM EDTA. Elution was achieved by incubating the beads with three bead volume of elution buffer for 15 min at 4°C on a rotator. Elution was repeated 3 times, the eluates were pooled and analyzed by mass spectrometry at SPARC BioCentre at the Hospital for Sick Children. Results were analyzed with Scaffold software (Proteome Software Inc.). Non-specific hits were eliminated with negative controls and comparisons with the Flag and EGFP mass spectrometry databases.

2.14 Expression and purification of NBEAL2 PH-BEACH domain

Bacterial constructs used for the expression of PH-BEACH domain of NBEAL2 are described in section 2.5.1. Constructs expressing 6XHis-PH-BEACH, 10XHis-PH-BEACH, 6XHis- NusA-PH-BEACH, 6XHis-MBP-PH-BEACH and 6XHis-Sumo-PH-BEACH were transformed independently into E.coli BL21(DE3), BL21(DE3) Star, and Rossetta (DE3). Source of the bacteria cultures used are described in section 2.3. The constructs were also transformed into BL21(DE3) Star bacteria that were pre-transformed with a combination molecular chaperone or disulfide isomerase expression plasmids to facilitate folding: pBB540 (GrpE, ClpB; addgene #27393), pBB542 (DnaK, DnaJ, higher expression of GroESL; addgene #27395), pBB550 (DnaK, DnaJ, lower expression of GroESL; addgene #27396), pBB572 (ibpA, ibpB; addgene #27397) and pBA2219 (DsbC; addgene #38152). Specifically, combinations of pBB540/pBB542/pBB572, pBB540/pBB550 and pBA2219 were used for the expression of PH-BEACH domain (de Marco, Deuerling et al. 2007, Austin and Waugh 2012). Transformed bacteria were grown in Luria-Bertani broth (LB broth; 1% w/v tryptone, 1% w/v NaCl, 0.5% w/v yeast extract, pH 7) or terrific broth (TB broth; 1.2% w/v tryptone, 2.4% w/v yeast extract, 0.5% w/v glycerol, 17 mM KH2PO4 and 72mM K2HPO4) supplemented with appropriate antibiotics until OD600 reaches 0.6. Protein expression was induced with 0.1-1mM of IPTG at a range of temperatures from 12°C to 37°C and at a range of time from 4h to overnight. Bacteria were pelleted (4000g, 20 min), resuspended in cold lysis buffer (50mM Tris, 300mM NaCl, 10mM imidazole, pH 8) and lysed with sonication following incubation with 1 mg/mL of lysozyme for 30 minutes on ice or with French press. Lysates were incubated with Ni-NTA resins (Qiagen) for 30min at 4°C on a rotator. The resins were then washed 3-5 times with wash buffer (50 mM Tris, 300 mM NaCl, 20 mM imidazole, pH 8) and eluted 3 times with elution buffer (50 mM Tris, 300 mM NaCl, 250 mM imidazole, pH 8). Resins were sometimes washed with ATP buffer (50 mM Tris, 5 mM ATP, 50 mM KCl and 20 mM MgCl2, pH8.0) following the imidazole washes. Eluates were pooled and further purified with gel filtration and/or cation exchange chromatography. Fraction from each step was collected for analysis to optimize the purification protocol.

PH-BEACH domain was expressed using the mammalian piggyBac expression system as previously described (Li, Michael et al. 2013). PH-BEACH domain of NBEAL2 was subcloned into PB-T-PAF vector as described in section 2.5.3. For transfection, 3X107 HEK293F cells were seeded into a shaking flask in 30 mL of FreeStyle 293 Expression Medium (Thermo Fisher). To prepare the transfection DNA-lipid complex, 28 μg of PB-T-PAF-NBEAL2, 3.5 μg

58 of PB-RB, and 3.5 μg of PBase plasmids from the piggyBac system and 35μL of FreeStyle MAX Reagent (Thermo Fisher) were mixed with 1.2 mL of OptiPRO SFM reduced serum medium (Thermo Fisher) and incubated at room temperature for 10min before addition to the HEK293F cell culture. 10 μg/mL of puromycin was added 24 h post-transfection to generate a stable cell line. For protein expression, HEK293F cells were resuspended in induction media (FreeStyle 293 Expression Medium containing 2 μg/mL of doxycycline (Sigma) for induction and 1 μg/mL of aprotinin as a protease inhibitor) at a density of 0.5X106 cells/mL. Cells were pelleted (300g, 5 minutes) on day 2 and resuspended in fresh induction media. Superntants containing the expressed proteins were collected and assayed with coomassie stains and western blot. Supernatants were collected every other day for 10 days and the cells were then discarded.

2.15 Fibrinogen endocytosis assays in mouse primary megakaryocytes

Mouse bone marrow cells were extracted by flushing mouse tibia and femurs with PBS, incubated in IMDM (Gibco) containing 10% BIT 9500 (Stemcell), 50 ng/mL rhTPO (gift from Kirin Brewery Company) and 1% penicillin-streptomycin (Wisent) for 3 days as described in section 2.3.2, and MKs purified on a 3%/1.5% BSA gradient as described in section 2.3.3. These MKs were either grown on Matrigel-coated coverslips and fixed on day 5, or kept in suspension culture for incubation with 0.1 mg/mL of FGN conjugated with Alexa Fluor 488 or 555 (Thermo Fisher). Incubation conditions for various experiments are described in the Figures and Results. After incubation cells were washed and centrifuged onto poly-lysine or matrigel coated coverslips (200g, 10 min). For FGN tracking experiments, day 5 MKs were incubated with 0.3 mg/mL of FGN-Alexa Fluor 488 for 30 minutes, then washed, seeded onto Matrigel coated coverslips and fixed at various time points (cells were spun onto coverslips for the t0 time point). All cells used for immunostaining were fixed with 4% paraformaldehyde (PFA; Electron Microscopy Sciences) in PBS.

2.16 Biotinylated fibrinogen for secretion and degradation assays

Human FGN (Haematologic Technologies) was biotinylated using EZ-Link Sulfo-NHS-Biotin (Thermo Fisher) according to the manufacturer’s instructions. To detect FGN secretion, day 3 purified MKs (above) were incubated with 0.1 mg/mL bio-FGN for 24 h; cells were then washed, filtered through a 100 µm mesh and put into fresh media, from which samples were collected at various time points and concentrated using centrifuge filters (Amicon) prior to SDS-PAGE immunoblot analysis (IB). Parallel cell samples were also lysed and analyzed. In

FGN degradation experiments MKs purified on day 4 incubated with 0.1 mg/mL of bio-FGN for 24 h; some cells were treated with chloroquine (50 µM) for 2 h prior to incubation. Cells were then washed, filtered, lysed and lysates analyzed via IB.

2.17 Flow cytometry and analysis

Endocytosed FGN was assessed by incubating day 4 mouse bone marrow culture cells with 0.1 mg/mL of Alexa Fluor 488 FGN conjugate overnight, or on day 5 for 1 h. Cells were then washed and labelled with rat anti-mouse CD41-PE (eBiosciences, 12041181, 1.25 µg/mL), filtered (100 µm mesh) to remove clumps analyzed via BD LSRII flow cytometer after the addition of 1 μg/mL of DAPI (Sigma) to detect non-viable cells and 0.1% trypan blue to quench surface-bound Alexa Fluor 488 FGN.

For MK ploidy analysis, cells from primary bone marrows and day 5 bone marrow cultures were centrifuged at 400g for 10 minutes, incubated with FITC conjugated anti-CD41 antibody (eBiosciences) at 4°C for 30 minutes and then fixed with 0.5% paraformaldehyde at room temperature for 20 minutes. Fixed cells were incubated with propidium iodide solution (50 μg/mL propidium iodide, 200 μg/mL RNase A, 0.1% Triton-X100) overnight and filtered through a 70 μm cell strainer. Flow cytometry was performed using BD FACSCanto (BD Biosciences) and the analysis was performed using FlowJo (Tree Star Inc.).

2.18 Imaging 2.18.1 Transmission electron and immunoelectron microscopy

Transmission electron microscopy (TEM) and immunoelectron microscopy (IEM) were performed by Richard Leung (Electron Microscope Technician).

For TEM, cells were fixed in 3% glutaraldehyde in 0.1 M cacodylate at pH 7.4, washed, post- fixed with 2% osmium tetroxide, dehydrated and embedded in Epon-Araldite. For IEM, cells were fixed in 0.5% glutaraldehyde in PBS pH 7.4 for 30 min at room temperature then 4-5h at 4°C. Cells were then washed with 0.1% glycine/PBS followed by 2% BSA/PBS prior to incubation (60 minutes) in primary antibody (rabbit anti-VWF or FGN). After repeated rinses with 2% BSA in PBS, cells were incubated (60 minutes) with goat-anti-rabbit IgG (H&L) conjugated with 10 nm gold (Electron Microscopy Sciences), rinsed with PBS and post-fixed with 2% osmium tetroxide in dH2O prior to dehydration and embedding in Epon-Araldite.

Thin sections were cut and stained with uranyl acetate and lead citrate, and samples were examined with a JEOL JEM-1011 electron microscope at 80 kV; images were captured with a side-mounted Advantage HR CCD camera (Advanced Microscopy Techniques, Danvers, MA) and processed with Adobe Photoshop.

2.18.2 Confocal immunofluorescence microscopy

Dilutions used for the primary antibodies are summarized in section 2.4.

Human primary and mouse bone marrow megakaryocytes were cultured as described in section 2.3.1 and 2.3.2 and seeded onto matrigel coated coverslips in 12-well plates. Cells were fixed with 4% paraformaldehyde for 30 minutes at room temperature, incubated with blocking buffer (PBS supplemented with 2% v/v horse serum and 2% w/v BSA) with or without 0.2% Triton X-100 for 1 h at room temperature. Cells were stained with primary antibodies for 1h at room temperature or overnight at 4°C. Cells were then washed with PBS (3 times, 5 minutes each wash) and stained with the corresponding fluorescent secondary antibodies for 1h at room temperature. Cells were then washed with PBS (3 times, 5 minutes each wash), stained with 1 μg/mL of DAPI (Sigma) for 10 minutes at room temperature, and mounted onto microscope slides with fluorescent mounting medium (DAKO) or prolong diamond (Thermo Fisher).

Cells were incubated with 0.1 mg/mL of mouse AffniPure Fab fragments (Jackson ImmunoResearch) for 1 h prior to incubation with primary antibodies when mouse cells were probed with mouse-derived antibodies. Fluorescent secondary antibodies used were diluted 1:1000 and obtained from Thermo-Fisher: donkey anti-mouse, rabbit, goat, sheep tagged with Alexa Fluor 488, 568 or 647; goat anti-rat Alexa Fluor 568. DAPI (Sigma) was used to stain nuclear DNA. For staining conditions involving two different rabbit antibodies, the antibodies were fluorescently labelled using the Zenon Alexa Fluor 555 Labelling Kit (Thermo Fisher) prior to staining.

Images were obtained with an oil immersion objective (60x, 1.35 N.A.) using a Quorum Olympus spinning-disc confocal inverted epifluorescence microscope equipped with four solid-state lasers (Spectral Applied Research): 405 nm, 491 nm, 561 nm and 642 nm, an Improvision Piezo Focus Drive, a 1.5x magnification lens (Spectral Applied Research), a

Hamamatsu C9100-13 back-thinned EM-CCD camera and a Yokogawa CSU X1 spinning- disc confocal scan head with the Spectral Aurora Borealis upgrade. Acquisition, deconvolution and image processing were performed using Volocity software (PerkinElmer). Figures were prepared with Adobe Photoshop.

2.19 Statistics

Statistical tests including Student’s t-test, analysis of variance (ANOVA), Mann-Whitney U test and Kruskal-Wallis test were performed using GraphPad Prism 6 (GraphPad Software) or SPSS version 25 (IBM).

Chapter 3

Investigating the molecular and structural properties of NBEAL2

3.1 Abstract

Subcellular fractionation experiments showed that NBEAL2 can adopt two distinct multimeric states in primary MKs: a lower molecular weight complex in the cytosolic fraction and a higher molecular weight complex distributed across cytosolic and membrane fractions. Subcellular localization analysis of NBEAL2 via immunofluorescence microscopy identified co-localization with P-selectin, an α-granule membrane marker. Co-immunoprecipitation experiments also showed interaction between NBEAL2 and P-selectin, supporting a functional role for NBEAL2 in α-granules. To explore the structural properties of NBEAL2, attempts were made to express and purify the PH-BEACH domain for X-ray crystallography analysis using a bacterial expression system. Success was limited, and a mammalian expression system offers new possibilities.

3.2 Introduction

It is predicted from in silico models that BDCPs can associate with vesicles to form scaffolds that enable interaction with other proteins, including BDCPs. Specific interactions predicted include WDFY3-LYST and WDFY3-NBEAL2 (Gunay-Aygun, Falik-Zaccai et al. 2011, Cullinane, Schaffer et al. 2013). BDCPs may also be capable of forming homodimers with functional significance. We examined the multimeric potential of NBEAL2 and its subcellular localization to better understand its properties and functions, and also endeavored to obtain information about its structure, specifically the PH-BEACH domain where numerous GPS- causing mutations reside (Fig. 1.10), via X-ray crystallography.

3.3 Results

Structured Illumination confocal microscopy imaging of platelets was performed by Dr. Fred Pluthero (Fig. 3.6 and Fig. 3.11A).

3.3.1 NBEAL2 forms oligomeric complexes

In silico models predict that BDCP potentially bind with each other (Cullinane, Schaffer et al. 2013), which raises the possibility that they may form homomultimers. To test whether NBEAL2 is capable of oligomerization, GFP-NBEAL2 and HA-NBEAL2 expression constructs were co-transfected into HEK293 cells. From these lysates, NBEAL2 was immunoprecipitated with either GFP or HA antibody, and the immunoblot was probed for the presence of the other

64 tagged NBEAL2 counterpart. The results indicate that GFP-NBEAL2 co-immunprecipitated with HA-NBEAL2 (Fig. 3.1A) and consistently, HA-NBEAL2 co-immunoprecipitated with GFP- NBEAL2 (Fig. 3.1B). These results suggest that NBEAL2 is capable of oligomerization in vitro.

Figure 3.1. NBEAL2 is capable of oligomerization. GFP- and Flag-NBEAL2 were co-transfected into HEK293 cells. The lysates were incubated with GFP or Flag antibody and the immunoprecitants were analyzed with immunoblots. (A) GFP-NBEAL2 co-immunoprecitated with HA-NBEAL2. (B) Reversing the pulldown condition, HA-NBEAL2 also immunoprecitated with GFP-NBEAL2. As HA-NBEAL2 and GFP-NBEAL2 are of similar molecular weights, pulldown efficiency was confirmed with Alexa Fluor 647 secondary to avoid residual signals from the previous HRP secondary. These results indicate that NBEAL2 is capable of forming dimers and/or higher order oligomers in vitro.

3.3.2 Validation of NBEAL2 antibody

Characterization of proteins is greatly aided by access to a valid specific antibody. After numerous attempts we were not able to generate a high quality anti-NBEAL2 rabbit antibody, but luckily antibodies became commercially available in the course of this study. After testing for specificity to endogenous NBEAL2 via immunoblotting we determined that the rabbit anti- NBEAL2 monoclonal antibody described in Material and Methods (Abcam) was suitable for our purposes. A single band of the expected molecular weight for NBEAL2 was detected in both normal human and WT mouse platelets, and found to be absent in both human GPS and Nbeal2-/- mouse platelets (Fig. 3.2).

Figure 3.2. Validation of NBEAL2 antibody and Nbeal2-/- mouse. Human GPS platelets and Nbeal2- /- mouse platelets were analyzed with NBEAL2 antibody. NBEAL2 expressions were absent in both human GPS platelets and Nbeal2-/- mouse platelets as expected. This validates the Nbeal2-/- mouse and the quality of NBEAL2 antibody in immunoblotting.

To test the suitability of this anti-NBEAL2 antibody for immunofluorescence microscopy, we attempted to generate a stable DAMI megakaryocytic cell line expressing tagged NBEAL2.

The idea was to verify the specificity of anti-NBEAL2 for immunolabelling by comparing the results to labelling for the tag.

An EGFP-NBEAL2-3XFlag mammalian expression construct was prepared and transfected into DAMI cells, but transfection efficiency appeared to be very low and no cells survived the subsequent puromycin antibiotic selection. Attempts to use lipid-mediated transfection (e.g. Lipofectamine 2000) and electroporation to achieve higher efficiency also failed. These failures are likely due to the large size of constructs containing NBEAL2 cDNA, which alone is 8.3kb. As an alternative approach, 3XFlag, EGFP and NBEAL2 were cloned into PLJM1 lentiviral vectors independently, and lentiviruses were prepared. Schematics of the constructs are shown in Fig. 3.3. Despite low transduction efficiency due to a large gene being packaged, lentivirus offers efficient gene integration once transduced, which is beneficial to the generation of a stable cell line. Following transduction, puromycin antibiotic selection, recovery and expansion, several clonal populations were obtained. Of the six DAMI cell populations, three appear to express the transgene (Fig. 3.4). Clonal population 2 was used for further experiments.

Figure 3.3. Schematic of NBEAL2 constructs used to generate stable cell lines. For routine expression of tagged NBEAL2, NBEAL2 was first subcloned into P3XFlag-CMV-14 vector and an EGFP fluorescent-tag was then added to its N-terminal (top panel). For lentiviral transduced expression of NBEAL2, NBEAL2 was subcloned into PLJM1 vector where 3XFlag and EGFP-tags were subsequently introduced to the N-terminal (lower panel).

Figure 3.4. Generation of DAMI cell lines stably expressing NBEAL2. DAMI cells were transduced with lentivirus packaged from PLJM1-3XFlag-EGFP-NBEAL2 and positive colonies were obtained with puromycin selection. Colonies were expanded and assayed for the expression of the transgene by probing the immunoblot with anti-Flag antibodies. The result confirmed the expression of 3XFlag- EGFP-NBEAL2 in several colony populations, specifically clonal population 2, 3 and 5.

To assess the quality of NBEAL2 antibody in immunofluorescence microscopy, we stimulated and fixed stable 3XFlag-EGFP-NBEAL2 DAMI cells, previously clonal population 2 as depicted in Fig. 3.4, and immunostained the cells with the NBEAL2 and Flag antibody. Pearson’s correlation coefficient (PCC) depicts a value of 0.914 ± 0.073 between NBEAL2 and Flag in these cells (Fig. 3.5, n=20 cells each for 3 independent experiments). Additionally, the NBEAL2 antibody does not stain human GPS platelets in contrast to control human platelets (Fig. 3.6). Altogether, these results indicate that the NBEAL2 antibody is adequate for both immunoblot and immunofluorescence analyses.

Figure 3.5. Validation of NBEAL2 antibody for immunofluorescence microscopy. Stable 3XFlag- EGFP-NBEAL2 DAMI cells were stimulated, fixed and stained with NBEAL2 and Flag antibody. Strong overlaps were observed between the NBEAL2 stains. Maturation of the cells did not impact the overlaps observed. This validates the suitability of the antibody for immunofluorescence. (Z-slice, scale bar = 5 µm).

Figure 3.6. Validation of anti-NBEAL2 antibody for immunostaining. Human platelets were immunostained for membrane-associated CD42B (green) and NBEAL2 (red), which were detected via appropriate fluorescent secondary antibodies. Imaging conditions were established to detect both proteins in normal donor platelets via confocal laser fluorescence microscopy (top), and the same settings were used to image platelets from a GPS patient (bottom) shown to be NBEAL2-null via immunoblotting with the same antibody. Non-deconvolved extended focus images captured with the same contrast and intensity settings show an absence of NBEAL2 signal in GPS platelets.

3.3.3 NBEAL2 forms a single molecular weight complex in DAMI cells

To further define the properties and nature of NBEAL2 and its complex, we explored the native conformation of the NBEAL2 complex in addition to determining whether it is cytosolic or membrane-associated. We utilized DAMI cells for further examination of NBEAL2. DAMI is a megakaryocytic cell line that normally express an abundant amount of endogenous NBEAL2. This approach was used to provide a more accurate depiction of NBEAL2’s properties.

We first prepared DAMI cell lysates as described in Fig. 3.7 where cytosolic and membrane fractions were obtained. Fractions were then loaded onto Blue Native and SDS-PAGE gels through a decreasing step gradient for visualization. In this context, Blue Native gels will provide information on the native conformation of NBEAL2 while SDS-PAGE gels will serve as loading controls. Resulting immunoblots were probed with NBEAL2 as well as CD61 antibody to ensure proper membrane loading. Interestingly, NBEAL2 was detected as a single molecular weight band at ~900 kDa (Fig. 3.8A). With previous results showing that NBEAL2 can at least form a dimeric structure and NBEAL2 being a 302 kDa protein, suggests that in DAMI cells, NBEAL2 in its native conformation forms a complex consisting of

70 a trimeric structure, or a dimeric structure with additional bound proteins. In both the Blue Native and SDS-PAGE gels, NBEAL2 was detected solely in the cytosolic fraction (Fig. 3.8A- B). This was unexpected since with their PH domains, BDCPs including NBEAL2 are predicted to be also membrane-associated.

Figure 3.7. Schematic for extracting cytosol and membrane fractions from cells. Cells were harvested and mechanically lysed by passing through a needle syringe aggressively against the bottom of a conical tube. Intact cells, debris and nuclear DNA were removed with centrifugation. Lysates were then subjected to ultracentrifugation for which the supernatant contains the cytosolic fraction and the pellet was reconstituted in Triton X-100 as the membrane fraction. The fractions were then analyzed with immunoblots.

Figure 3.8. NBEAL2 is primarily cytosolic in DAMI cells. DAMI cells were fractionated into cytosolic and membrane fractions and the presence of NBEAL2 was analyzed with immunblots with decreasing sample loading across the lanes. (A) DAMI cytosolic and membrane fractions were loaded onto Blue Native gel, transferred onto PVDF membrane and probed with NBEAL2. (B) DAMI cytosolic and membrane fractions were denatured and reduced prior to loading onto SDS PAGE gel. The proteins were then transferred onto nitrocellulose membrane and probed with NBEAL2. CD61 controls for the membrane loading. In both conditions, NBEAL2 is predominantly cytosolic.

3.3.4 Two distinct NBEAL2 complexes in primary human MKs are both cytosolic and membrane-associated

With NBEAL2 being observed as a single cytosolic ~900 kDa complex in DAMI cells, we then ask if these properties of NBEAL2 are consistent in other megakaryocytic cell types. While DAMI cell lines are megakaryocytic cells, they do not form recognizable α-granules. This raises caveats when using this cell line to assess NBEAL2 function, considering NBEAL2 is essential for α-granule biogenesis in humans. To address this issue, primary human MKs were utilized. Cultured primary MKs from human CD34+ cells were lysed and fractionated as described in Fig. 3.7. Total lysate, cytosolic and membrane fractions were loaded onto Blue Native and SDS-PAGE gels for analysis. The fractions were loaded in a balanced manner such that the amount of contents of cytosolic and membrane fractions loaded constitute the amount of contents of the total lysate loaded. In this manner, different fractions can be directly compared. From the Blue Native immunoblots as shown in Fig. 3.9, NBEAL2 was detected as two distinct molecular weight complexes, ~600 kDa and ~900 kDa. This is in contrast to the single ~900 kDa NBEAL2 complex found in DAMI cells. Interestingly, the ~600 kDa NBEAL2 complex is more prevalent in the cytosolic fraction whereas the ~900k Da complex is distributed equally between the cytosolic and membrane fractions. Densitometric analysis of the SDS-PAGE immunoblot revealed that 66.9±2.1% of NBEAL2 is cytosolic and 36.0±1.8% of NBEAL2 are membrane-associated. As expected in both the Blue Native and SDS-PAGE immunoblots, P-selectin, VWF and SEC22B are membrane-associated whereas GAPDH is predominately cytosolic. These results reveal important understanding on the nature of the NBEAL2 complexes in primary human MKs.

Figure 3.9. Fractionation of primary human megakaryocytes. Lysates derived from mechanical cell lysis of primary human MKs were first centrifuged at 3,000g for 10 minutes to remove cell debris and DNA. Lysate was then ultracentrifuged at 120,000g for 2hrs. Supernatant was collected as the cytosolic fraction and the pellet was solubilized in Triton X-100 and collected as the membrane fraction. Blue Native and SDS-PAGE of the total lysate, cytosolic and membrane fractions were immunoblotted for NBEAL2, SEC22B, P-Selectin, VWF and GAPDH. NBEAL2 is detected in both the cytosolic and membrane fractions as two distinct complexes with the lower molecular weight configuration been more prevalent in the cytosolic fraction.

3.3.5 NBEAL2 localizes primarily to the α-granule marker, P-selectin, in human primary MKs

Understanding the subcellular localization of NBEAL2 will offer insights into its function. With a validated NBEAL2 antibody, we are able to study NBEAL2’s subcellular localization within primary human MKs. It is especially important to study primary MKs with regards to NBEAL2 since most available megakaryocytic cell lines contain no discernable α-granules when examined by TEM. Since NBEAL2 is essential in α-granule biogenesis, these cell lines may not provide an accurate depiction of the subcellular localization of NBEAL2. For localization analysis, primary human MKs were immunostained with NBEAL2 against an array of cellular organelle markers: ER (calnexin), cis-Golgi (GM130), trans-Golgi (TGN46), trans-Golgi vesicles (AP1), α-granule membrane (P-selectin), δ-granules (CD63), plasma membrane protein (TFR), early endosomes (RAB5 and EEA1), late endosomes (RAB7), recycling endosomes (RAB11) and α-granule cargoes (VWF and TSP1). Representative confocal microscopy images are shown in Fig. 3.10A. Manders overlap analysis of NBEAL2 against these markers revealed the highest score with P-selectin with an overlap coefficient of 0.753േ0.161 (Fig. 3.10B). Other cellular compartment markers do not colocalize strongly with NBEAL2 (Manders coefficient <0.4). These results indicate that in primary human MKs, NBEAL2 is most likely found with the α-granule membranes, suggesting a role of NBEAL2 directly at the α-granules.

Figure 3.10. NBEAL2 colocalizes with P-selectin in primary human megakaryocytes. (A) Representative immunofluorescence microscopy images of cultured human MKs stained for NBEAL2 and various cellular markers (Z-slice; scale bar = 5 µm). (B) Manders overlap coefficient analysis showing prominent NBEAL2 colocalization with P-selectin (n=3 independent experiments with 15-20 cells for each condition in each experiment).

3.3.6 NBEAL2 interacts with P-selectin

Previous co-localization analysis of NBEAL2 in primary human MKs has revealed a strong co-localization between NBEAL2 and P-selectin, an α-granule membrane marker. This prompts the question of whether NBEAL2 could potentially interact with P-selectin. The results could yield important knowledge on the molecular mechanism of action of NBEAL2. NBEAL2 is observed to co-localize with P-selectin in human platelets similar to that seen in human primary MKs (Fig. 3.11A). To test interaction, NBEAL2 or P-selectin was immunoprecipitated from human platelet lysates then immunoblotted for P-selectin or NBEAL2 respectively to detect potential interaction. The results revealed an interaction between NBEAL2 and P-selectin (Fig. 3.11B). Similar results were seen when primary human MKs were used (Fig. 3.12). Altogether, these results indicate that NBEAL2 and P- selectin interacts, further suggesting a role for NBEAL2 at the α-granules.

Figure 3.11. NBEAL2 interacts with P-selectin. (A) Substantial overlap was observed between NBEAL2 and P-selectin in co-stained human platelets (3D renders plus overlap channel generated from structured illumination microscopy images; bar = 0.8 µm). (B) Co-immunoprecipitation analysis using human platelet lysates. NBEAL2 or P-selectin were immunoprecipitated from human platelet lysates, and the co-immunoprecipitants were analyzed by immunoblot. P-selectin co- immunoprecipitated with NBEAL2 (left panel) and NBEAL2 co-immunoprecipitated with P-selectin (right panel). These results indicate interaction of NBEAL2 with P-selectin within platelets.

Figure 3.12. NBEAL2 interacts with P-selectin in cultured primary human megakaryocytes. Day 12 cultured human primary MKs were lysed, P-selectin was immunoprecitated and analyzed with immunoblot. The results reveal native interactions between NBEAL2 and P-selectin in primary MKs.

3.3.7 Expression and purification of the PH-BEACH domain of NBEAL2

PH-BEACH domain defines the BDCP family. As shown in Fig 3.13A, numerous missense mutations known to cause GPS are also found in the PH-BEACH domain of NBEAL2. It is thus of interest to understand the structural properties of this domain and of the critical residues that are essential to NBEAL2’s function. To visualize the structural properties of NBEAL2’s PH-BEACH domain, x-ray crystallography or cryo electron microscopy need to be employed. To prepare the proteins, PH-BEACH domain of NBEAL2 (1918aa-2348aa) was subcloned into pET28a bacterial expression vector then transformed into bacterial host strain BL21. Expression of the recombinant NBEAL2 can be controlled via T7 promoter that is inducible by IPTG. The recombinant protein contains a 6XHis tag at its N-terminal to enable purification from the crude bacterial lysate as shown in Fig 3.13B.

Transformed BL21 bacteria were induced with IPTG for 4 h, 6 h or overnight at 16°C, 25°C or 37°C. Cells were then lysed with sonication or French press to better preserve the native conformation of the protein. Insoluble materials were sedimented with centrifugation at 3000g for 10 min and the resulting pellet was solubilized with Triton X-100. Soluble crude lysates

79 were then loaded onto Ni-NTA column to facilitate binding. The column was washed up to 5 times with wash buffer containing imidazole with concentration ranging from 20 mM to 50 mM. 6XHis-PH-BEACH was then eluted from the column with 250 mM imidazole. Samples from various steps of the purification were examined with coomassie blue gels.

Despite altering the IPTG concentration, induction time and induction temperature, similar purification results were obtained. A representative Coomassie blue stained gel of a typical purification is shown in Fig. 3.14. Transformed BL21 bacteria were grown to OD600=0.4, then induced with 0.2 mM of IPTG overnight at 16°C. Three distinct protein bands near 63 kDa were observed. From the highest to the lowest molecular weight, the identities of these proteins were identified with mass spectrometry to be DnaK, GroEL and NBEAL2 PH-BEACH. In attempt to remove DnaK and GroEL chaperones from the PH-BEACH protein, beads were additionally washed with ATP buffer following the imidazole washes. Unfortunately, this approach was of limited success as majority of DnaK and GroEL were still present in the elution despite repeated ATP washes.

Figure 3.13. Schematic of the NBEAL2 PH-BEACH construct. (A) Full-length NBEAL2 contains numerous domains and missense mutations (marked in red) known to cause GPS reside in these domains. (B) PH-BEACH domain of NBEAL2 (1918-2345aa) was subcloned into bacterial protein expression. 6X or 10X His-tags were used in combination with nickel beads to affinity purify the recombinant proteins.

Figure 3.14. Expression and purification of PH-BEACH. 6XHis-PH-BEACH domain of NBEAL2 (1918-2345aa, molecular weight 54 kDa) was expressed in bacterial strain BL2. Bacteria were grown to OD600=0.4, induced with 0.2 mM of IPTG overnight at 16°C. Lysates were incubated with nickel Ni- NTA beads and washed. PH-BEACH proteins were eluted with the indicated imidazole concentration. Fractions from different steps of the purification were analyzed with coomassie blue gels. Majority of the recombinant PH-BEACH protein was insoluble. Two major contaminating proteins were co-eluted with the soluble PH-BEACH protein and were identified by mass spectrometry as DnaK and GroEL.

To obtain a higher purity of PH-BEACH protein for structural studies, secondary purifications were employed. PH-BEACH proteins were eluted from the nickel bead column as described in Fig 3.14 then subjected to cation exchange chromatography. Proteins were eluted from the column through a NaCl gradient. Collected fractions were then analyzed by Coomassie blue stains. As shown in Fig. 3.15, PH-BEACH protein was eluted at a NaCl concentration of 622 mM. The fraction was however still heavily contaminated with DnaK (contam. 1) and GroEL (contam. 2). As an alternative approach, gel filtration was utilized following nickel bead purification. The majority of the PH-BEACH protein was found in fractions 15-17 (Fig. 3.16). Though similar to the results observed with cation exchange chromatography, these fractions were still contaminated with DnaK and GroEL. Taken together these results suggest that DnaK and GroEL are tightly bound to PH-BEACH protein and are difficult to separate. Alternative approaches are required to remove these contaminants.

Figure 3.15. Purification of PH-BEACH with cation exchange chromatography. PH-BEACH domain of NBEAL2 (1918-2345aa, molecular weight 54kDa) was purified with nickel column then subjected to cation exchange chromatography (HiTrap SP HP). Proteins were eluted on a linear NaCl gradient. Collected fractions were then analyzed with coomassie gels. The elution peak of PH-BEACH protein overlaps significantly with contamination protein 1 (Contam. 1), later identified as DnaK, and contamination protein 2 (Contam. 2), later identified as GroEL.

Figure 3.16. Purification of PH-BEACH with gel-filtration chromatography. PH-BEACH domain of NBEAL2 (1918-2345aa, molecular weight 54kDa) was purified with nickel column then subjected to gel-filtration chromatography (Sephadex G-100). Fractions were collected as proteins pass through the column and the fractions were then analyzed with coomassie gels. The elution peak of PH-BEACH protein overlaps significantly with contamination protein 1 (Contam. 1) and contamination protein 2 (Contam. 2). Contam. 1 and contam. 2 appeared to be DnaK and GroEL but are indistinguishable due to significant peak overlaps. Vo indicates the void volume.

Several key observations suggest that the PH-BEACH protein is likely misfolded in this bacterial expression system. Firstly, most of the PH-BEACH protein expressed are insoluble (Fig. 3.14), suggesting the formation of inclusion bodies or insoluble aggregates. Secondly, the detection of DnaK and GroEL chaperones that are inseparable from the PH-BEACH protein indicates the presence of unstructured protein. In light of these findings, attempts were made to improve the solubility and folding of PH-BEACH proteins. Two main strategies were employed to accomplish this. Firstly, tags that are known to improve solubility of recombinant proteins were engineered into the PH-BEACH protein. These tags include NusA, MBP and Sumo (Fig. 3.17A) (Marblestone, Edavettal et al. 2006, Costa, Almeida et al. 2014). 10XHis-PH-BEACH was designed to enable higher stringency washings to hypothetically remove more contaminants during the purification process. Secondly, since it is not unusual

83 for recombinant mammalian proteins to be poorly folded in bacteria due to inefficiencies and thus leading to the formation of inclusion bodies, a common approach is to co-express bacterial folding machineries to facilitate this process. To improve the expression and foldability of PH-BEACH proteins, several combinations of bacterial chaperones and protein disulfide isomerases were co-expressed alongside with the PH-BEACH protein. As suggested by (de Marco, Deuerling et al. 2007, Austin and Waugh 2012), specific bacterial chaperone combinations were used: pBB540 (GrpE, ClpB)/pBB542 (DnaK,DnaJ, higher expression of GroESL)/pBB572 (ibpA, ibpB), pBB540 (GrpE, ClpB)/pBB550 (DnaK, DnaJ, lower expression of GroESL) and pBA2219 (DsbC). Rossetta is an E.coli BL21 bacterial strain containing plasmids encoding for rare tRNA whereas BL21 Star is a BL21 bacterial strain that offers higher protein expression due to the inactivation of RnaseE (rne131) and outer membrane (OmpT) proteases (Sorensen and Mortensen 2005). Schematics of constructs and bacterial hosts used are shown in Fig 3.17A-B. Tagged PH-BEACH proteins were transformed into these bacterial strains and purified as described previously. Despite these efforts, similar results to Fig. 3.14 were observed in all the conditions tested, where DnaK and GroEL remain persistent in the elution fraction. New approaches are required to overcome this challenge.

Figure 3.17. Strategies to improve PH-BEACH protein quality. (A) PH-BEACH domain was tagged with 6XHis, 10XHis, NusA, MBP or Sumo. The aim is either to enable a more stringent purification condition or to improve protein solubility. (B) To encourage proper protein folding, multiple strains of BL21 bacteria were generated and used to express PH-BEACH. Rossetta strain harbours plasmids encoding rare tRNA codons in bacteria. BL21 Star strain generates higher amount of soluble proteins. pBB540/542/550/572 and pBA2219 plasmids encode for bacterial chaperons and protein disulfide isomerases with the aims to improve protein foldability and solubility.

While many recombinant mammalian proteins can be successfully expressed and purified from E.coli bacteria, many mammalian proteins can not. For these proteins, a rational is that E.coli simply lacks the machineries that are required for the proper folding of mammalian proteins (Fakruddin, Mohammad Mazumdar et al. 2013). To overcome the challenge of improperly folded PH-BEACH protein in the bacterial expression system, a mammalian expression system was used. Using the piggyBac expression system, PH-BEACH was subcloned into PF-T-PAF vector and expressed in suspension HEK293F and adherent HEK293T cells (Li, Michael et al. 2013). Culture supernatants were then collected for up to 10 days and visualized with immunoblot. An initial screen indicated the successful expression of the PH-BEACH proteins from these cells (Fig. 3.18). Further improvements and modifications are in progress to obtain higher quality proteins.

Figure 3.18. Expression of PH-BEACH in mammalian cells. PH-BEACH domain was expressed using the piggyBac system. PH-BEACH was subcloned into PB-T-PAF and expressed in HEK293T or HEK293F cells. Day 4 culture supernatants were collected and analyzed with immunoblot. ProteinA- tagged PH-BEACH was detected at the correct predicted molecular weight.

3.4 Discussion 3.4.1 NBEAL2 and PH-BEACH family proteins as oligomeric complexes

In our study, we have found that NBEAL2 is capable of forming oligomers (Fig. 3.1). This result is perhaps not unexpected as it was previously predicted that members of BDCP may cross-interact, however this is an untested theory where the potential of oligomerization of individual BDCP has not been examined (Albers, Cvejic et al. 2011, Cullinane, Schaffer et al. 2013). Since members of BDCP share a high degree of similarity in protein architecture (Fig. 1.11), it is plausible that if BDCP can indeed cross-interact that they may also be able to homodimerize. An unanswered question is how oligomerization influences NBEAL2’s function.

Typically, protein oligomerization does not occur through coincidence and the formation of oligomers such as homodimers are essential and of critical importance to the function of a protein. Such properties may contribute to the stability of the protein, offer functional gains, or act as a regulatory mechanism (Fakruddin, Mohammad Mazumdar et al. 2013). Defects in protein oligomerization often leads to protein dysfunction and are commonly associated with diseased states. This is the case with certain types of VWF disease where defects in VWF multimerization results in a severe bleeding disorder (Tjernberg, Vos et al. 2004). Acknowledging these factors, a plausible hypothesis is that NBEAL2 oligomerization is functionally important and defects in oligomerization may account for some cases of GPS. An initial approach to test this hypothesis may be to determine if any of the known GPS-causing mutations abrogates the oligomerization properties of NBEAL2.

3.4.2 NBEAL2 displays variable properties in DAMI cells and in primary human MKs

Blue-Native PAGE analyses have shown that NBEAL2 appeared as a single molecular weight complex (~900 kDa) and as two distinct molecular weight complexes (~600 kDa and ~900 kDa) when DAMI cell lysates and primary MK lysates were tested respectively (Fig. 3.8 and Fig. 3.9). While the 900 kDa NBEAL2 complex in both cell types may be of similar nature, the appearance of an additional 600 kDa NBEAL2 complex in primary human MKs is puzzling. Further, the 900 kDa NBEAL2 complex seen in DAMI cells is only present in the cytosolic fraction but not in the membrane fraction, while both the 600 kDa and 900 kDa NBEAL2 complexes are present in both the cytosolic and membrane fraction of primary MKs. These discrepancies suggest there are fundamental differences in the action and complexing properties of NBEAL2 in these cell types.

It is understood that with the exception of a recently introduced imMKCL cells, most megakaryocytic cell line models are incapable of forming proper α-granules (Nakamura, Takayama et al. 2014). Unfortunately, this includes DAMI cells which we have examined via TEM but were unable to detect structurally distinct α-granules. Inevitably, this leaves primary MKs as the preferred cell type to study α-granule biogenesis. Acknowledging these factors, it is reasonable that if NBEAL2 resides primarily near α-granules, as suggested by the results shown in Fig. 3.10, Fig. 3.11 and Fig. 3.12, its functional and complexing properties are more likely to be accurately represented by primary MKs, that generate true α-granules, rather than by DAMI cells. The absence of 900 kDa NBEAL2 complex in the membrane fraction of DAMI

87 cells also supports this view. It should be noted that for DAMI cells, while NBEAL2 complex was not detected in the Blue-Native gel, NBEAL2 was present in membrane fractions when analyzed by denaturing SDS-PAGE gels (not shown), suggesting that membrane-associated NBEAL2 in DAMI cells may represent an abnormal complex association. Considering that true α-granule formation is not observed in DAMI cells, may indicate the importance of functional NBEAL2 complex formation in α-granule biogenesis.

The discrepancies among the cell types also demonstrate some of the difficulties and challenges involved in studying α-granule biogenesis. Few appropriate alternative cell models are available besides primary MKs. Nevertheless, the presence of 600 kDa and 900 kDa NBEAL2-containing complexes in primary MKs suggest that NBEAL2 may exist in two conformations in the cell. Since NBEAL2 is a 302 kDa protein, suggests the possible presence of a homo-dimer in the 600 kDa complex and a homo-trimer or homo-dimer with additional interacting proteins in the 900 kDa complex. It is also possible that the differences in protein structure conformation may have resulted in these observations (e.g. elongated vs rod-shaped proteins). Whether these NBEAL2 complexes have different functional properties needs to be explored further.

3.4.3 Distinct configuration states of NBEAL2 in the cytosol and with membranes

Blue-Native gel analysis has revealed that NBEAL2 forms two distinct complexes with different molecular weights in primary MKs (Fig. 3.9). These complexes were detected in both the cytosolic and membrane fractions and are therefore not mutually exclusive. The MK fractionation results suggest that the 900 kDa NBEAL2 complex is approximately equivalent in the cytosol and membrane, whereas the 600 kDa NBEAL2 complex appears higher in the cytosolic fraction compared to the membrane fraction. While the implications of this phenomenon are unclear, these results provide some insights to the subcellular localization of NBEAL2. My results are the first demonstration showing that NBEAL2 may reside both in the cytosol and membrane within human MKs where it may have distinct functions during α- granule biogenesis.

Since the α-granule cargo proteins VWF and TSP1, were detected only in the membrane fraction suggests that α-granules remained intact during the fractionation process, eliminating the possibility of α-granule lumen proteins leaking into the cytosol. The exclusive presence of

SEC22B and P-selectin, an ER membrane protein and α-granule membrane protein respectively, in the membrane fraction also confirms that there were no contaminating membranes in the cytosolic fraction. If NBEAL2 is primarily an α-granule luminal or α-granule membrane protein, it should have been primarily detected in the membrane fraction together with VWF and P-selectin (Fig. 3.8). However since the majority of NBEAL2 was found in the cytosolic fraction, suggests that NBEAL2 is a cytosolic protein that acts on the α-granules from the cytosolic interface. It is possible that NBEAL2 functions by tethering to the α-granule membrane, a hypothesis that is supported by the interaction between NBEAL2 and P-selectin (Fig. 3.11 and Fig. 3.12), hence a fraction of NBEAL2 is observed in the membrane fraction.

3.4.4 NBEAL2 functions at α-granules

Immunofluorescence microscopy revealed that NBEAL2 localizes to P-selectin in primary MKs (Fig. 3.10), a result that is supported by the interaction of NBEAL2 with P-selectin and immunogold labelling in MKs (not shown). While it cannot be excluded that NBEAL2 may be able to perform other cellular functions, as NBEAL2 does partially co-localize with other cellular compartment markers, it is nevertheless clear that NBEAL2 co-localizes with P- selectin most significantly. This suggests that NBEAL2 has a functional role at the α-granules. In conjunction with the evidence that a large proportion of NBEAL2 is cytosolic and a smaller proportion of NBEAL2 is membrane-associated, one may surmise that NBEAL2 tethers to α- granule membranes from the cytosolic side and acts specifically at the α-granules. NBEAL2 is well positioned to act as a scaffold protein, the role predicted for BDCP, and it has the potential to regulate α-granule biogenesis through various means, including but not limited to the facilitation of membrane trafficking events and the formation of MCS. Both of these hypothetical functions of NBEAL2 could potentially influence the development and maturation of α-granules and may potentially offer an explanation to the phenotypes seen in GPS. More detailed functions of NBEAL2 at the α-granules will be explored in a later chapter.

3.4.5 PH-BEACH domain may direct the localization of NBEAL2

We have shown that NBEAL2 resides at the α-granules. However, it is unclear how this occurs. One possibility is that NBEAL2 binds to P-selectin directly. Sequence analysis predicts that a large proportion of P-selectin actually faces inwards towards the α-granule lumen, leaving only a small C-terminal tail, consisting of 35 residual amino acids, for

89 interaction at the cytoplasmic side (Johnston, Cook et al. 1989). While it is possible that NBEAL2 may interact with P-selectin via this cytoplasmic tail, an alternative prediction is that NBEAL2 interacts with P-selectin indirectly. One candidate is the interaction between NBEAL2’s PH-BEACH domain with the phospholipid constituents of the α-granule membrane. Previous studies have revealed that there are functional variabilities among the PH-BEACH domains of BDCP. For example, PH-BEACH domain from NBEA does not bind to phospholipids whereas the PH-BEACH domain from NSMAF does (Gebauer, Li et al. 2004, Haubert, Gharib et al. 2007). Further, for the BDCP that were confirmed to bind to phospholipids, specificity and selectivity were observed: WDFY3 specifically binds to PI3P and NSMAS selectively binds to PI4,5P (Simonsen, Birkeland et al. 2004, Haubert, Gharib et al. 2007). These studies suggest that there exists a potential that NBEAL2 may bind to specific phospholipid constituents on α-granules via its PH-BEACH domain. If this is true, then it is likely that the PH-BEACH domain of NBEAL2 directs the association of NBEAL2 with α-granules. Further investigations will be required to test this hypothesis.

3.4.6 GPS mutations in the PH-BEACH domain provide mechanistic clues

Possibly due to the lack of proper folding machineries in bacterial cells, we were not able to obtain PH-BEACH proteins of sufficient quality to proceed with structural elucidation. The crystal structures of PH-BEACH domain from other members of the BDCP family, NBEA and LRBA, have been previously elucidated (Jogl, Shen et al. 2002, Gebauer, Li et al. 2004). Based on these crystal structures, it was proposed that the PH and BEACH domain are likely functioning as a single entity rather than being independent domains. The structure of the PH-BEACH domain of NBEAL2 was constructed using Phyre2 protein structure modelling using the NBEA PH-BEACH as the modelling template (Kelley, Mezulis et al. 2015). By this modelling, it is revealed that all six of the identified GPS-causing missense mutations in the PH-BEACH domain of NBEAL2 (R2071P, P2100L, H2159Q, H2263Y, S2269L and R2345W) are positioned in close proximity and appear to surround a single empty cleft. This indicates the likely presence of an active site whereby when these amino acids are mutated, the function of this active site is lost, leading to the dysfunction of NBEAL2. Despite being a structural prediction, it nevertheless hints to a mechanism of the PH-BEACH domain which has not previously been studied. The validity of this hypothesis requires the elucidation of the precise structure of NBEAL2’s PH-BEACH domain. To this end, the expression of the PH- BEACH domain in a mammalian system shows promise for future structural studies (Fig. 3.18).

Chapter 4

Identification and characterization of the binding partners of NBEAL2

4.1 Abstract

NBEAL2 is essential for the formation of α-granules in MKs and platelets, but the mechanisms involved are unknown. To begin to understand the mechanism and function of NBEAL2, binding partners of NBEAL2 in DAMI cells were explored. Among the potential interacting partners identified via immunoprecipitation mass spectrometry, 14-3-3, FLNA and SEC22B were confirmed as genuine interactors of NBEAL2. Binding region delineation revealed that 1798-1903aa of NBEAL2 binds to SEC22B. Two missense mutations reported to cause GPS, E1833K and R1839C, reside within this region. NBEAL2 harboring either of these mutations fails to bind to SEC22B. These results suggest that the interaction between NBEAL2 and SEC22B may be critical for α-granule biogenesis.

4.2 Introduction

Prior to its link to GPS, NBEAL2 was a protein of unknown role and function. BDCP have been proposed to function as scaffold proteins (Cullinane, Schaffer et al. 2013), and if it is also a scaffold, NBEAL2 should interact with other proteins. Identification of interacting partner(s) with known function should provide insights into NBEAL2 function.

4.3 Results 4.3.1 Identification of NBEAL2 binding partners using yeast-two-hybrid screens We employed a yeast two-hybrid system to screen for potential NBEAL2 binding proteins. Since yeast cells express large proteins like NBEAL2 poorly, constructs expressing N- terminal and C-terminal portions of NBEAL2 were prepared as shown in Fig. 4.1, then subcloned into pGBKT7 yeast expression vector. The truncated NBEAL2 constructs were transformed into AH109 yeast cells as bait, and expression was confirmed via immunoblotting (Fig. 4.2). Transformed AH109 cells were subsequently mated with Y187 yeast cells that were pre-transformed with human bone marrow cDNA library as prey. Since NBEAL2 is primarily expressed in bone marrow haematopoetic cells, this approach was expected to provide more relevant results than using a universal cDNA library (Albers, Cvejic et al. 2011). Prey proteins from the positive colonies obtained from x-α-galactosidase plates were sequenced and identified with NCBI BLAST (Basic Local Alignment Search Tool). Results are listed in Table 4.1 and Table 4.2 for N-terminal and C-terminal NBEAL2 constructs respectively. Common contaminants including ferritin and hemoglobin were removed from the lists.

Figure 4.1. Schematic of NBEAL2 constructs used for the yeast-two-hybrid screens. N-terminal (1-1470aa) and C-terminal (1404-2754aa) of NBEAL2 were subcloned into pGBKT7 vector as bait proteins for yeast-two-hybrid. This approach will approximate the interacting regions of NBEAL2 with the hits obtained.

Figure 4.2. Expression of NBEAL2 bait constructs. N- and C-terminal NBEAL2 bait constructs were transfected into AH109 yeast and the expressions were analyzed with immunoblots using myc antibody. Both N- and C-terminal NBEAL2 were detected at the correct molecular weights of approximately 150kDa.

Table 4.1. List of potential NBEAL2 N-terminal interaction partners from yeast-two- hybrid screens. NBEAL2 N-terminal (1-1470aa) Gene Hits CD59 molecule 1 chromosome X open reading frame 61 1 COP9 signalosome subunit 6 (COPS6) 1 cyclin-dependent kinase inhibitor 3 1 EF-hand calcium binding domain 10, RAD50 interactor 1 1 eukaryotic translation initiation factor 3, subunit F 2 IKAROS family zinc finger 5 1 isocitrate dehydrogenase 2 1 lectin, galactoside-binding, soluble, 1 2 major histocompatibility complex 1 McKusick-Kaufman syndrome 1 mitochondrial translational release factor 1-like 1 natural killer cell group 7 sequence 1 PNN-interacting serine/arginine-rich protein 1 proteasome assembly chaperone 2 1 RPTOR independent companion of MTOR 1 serine carboxypeptidase 1 1 serine/arginine-rich splicing factor 9 2 signal recognition particle 68kDa 1 STAM binding protein-like 1 1 synaptonemal complex protein 1 1 U3 small nucleolar RNA-associated protein 6 homolog 1 zinc finger RNA binding protein 1

Table 4.2. List of potential NBEAL2 C-terminal interaction partners from yeast-two- hybrid screens. NBEAL2 C-terminal (1404-2724aa) Gene Hits actin, Homo sapiens POTE ankyrin domain family 4 adaptor-related protein complex 1, beta 1 subunit (AP1B1) 2 aldolase A, fructose-bisphosphate 1 amyloid beta precursor protein-binding, family B, member 2 1 casein kinase 2 1 CDK5 regulatory subunit associated protein 3 1 chromosome 17 open reading frame 62 (C17orf62) 1 chromosome 6 open reading frame 108 1 COP9 signalosome subunit 6 (COPS6) 1 DAZ associated protein 2 (DAZAP2) 3 heterogeneous nuclear ribonucleoprotein U-like 1 1 HtrA serine peptidase 2 1 major histocompatibility complex 2 Mannosidase 2 matrix metallopeptidase 25 2 NADH dehydrogenase (ubiquinone) 1 beta subcomplex 1 NADH dehydrogenase Fe-S protein 2 1 plakophilin 4 (PKP4) 1 poly (ADP-ribose) polymerase family 1 proline-rich transmembrane protein 4 (PRRT4) 1 Prosaposin 1 RNA (guanine-7-) methyltransferase 1 superkiller viralicidic activity 2-like 2 1 translocator protein (18kDa) (TSPO) 1 tumor protein p63 regulated 1-like 1 ubiquitin specific peptidase 22 1

While numerous hits were obtained from the yeast-two-hybrid assay, most protein hits do not share a common function, making it difficult to determine their significance to the function of NBEAL2. As we have previously hypothesized that NBEAL2 is likely involved in α-granule protein trafficking, we examined protein hits that have known functions in vesicular trafficking. These proteins include COP9 signalosome subunit 6 (COPS6) for both N- and C-terminal NBEAL2 and specifically for C-terminal NBEAL2, adaptor-related protein complex 1 (AP1) and DAZ associated protein 2 (DAZAP2). To confirm these interactions with NBEAL2, co- immunoprecipitation assays were conducted. GFP-NBEAL2 constructs were transfected into HEK293 cells, GFP-NBEAL2 was then immunoprecipitated from the lysate and examined via immunoblot. Based on these co-immunoprecipitation results, interactions between NBEAL2 and AP1, DAZAP2 or COPS6 were negative. With most of the other protein hits sharing no commonalities and having no relations to protein trafficking, we decided to utilize alternative approaches in finding NBEAL2’s binding partners.

Figure 4.3. Confirmation of yeast-two-hybrid hits. 3XFlag-EGFP-NBEAL2 was transfected into HEK293 cells, NBEAL2 was immunoprecipitated from the resulting lysate and the co- immunoprecipitants were analyzed with immunoblots using AP1, DAZAP2 or COPS6 antibodies. Results did not indicate that NBEAL2 interacts with these proteins.

4.3.2 Identification of NBEAL2 binding partners in DAMI cells using affinity purification mass spectrometry

Since yeast two-hybrid screens did not provide us with satisfactory results, we decided to pursue an alternative approach to identify NBEAL2 binding partners that employed megakaryocytic DAMI cells. DAMI cells expressing 3XFlag-EGFP-NBEAL2 described in Fig. 3.4, Fig. 3.5 and Fig. 3.6 were utilized for this purpose. NBEAL2 was affinity purified from cell

96 lysates with mouse monoclonal anti-Flag antibody and the eluates were subsequently analyzed via mass spectrometry. Half of the lysate volume was incubated with mouse IgG beads to exclude non-specific interactions. Experiments were repeated three times and the results were filtered for the most consistent hits. Results of interest are shown in Table 4.3.

Table 4.3. Potential NBEAL2 binding partners obtained from 3XFlag-NBEAL2 IP mass spectrometry with DAMI cells. Mass Spectrometry Results 14-3-3 epsilon 14-3-3 gamma 14-3-3 zeta 14-3-3 theta 14-3-3 beta 14-3-3 eta Filamin A (FLNA) Vesicle-trafficking protein SEC22b (SEC22B) 150+ other proteins

4.3.3 Verification of NBEAL2 binding partners

14-3-3, FLNA and SEC22B were consistently detected as NBEAL2 interactors in repeated experiments. Filamin A (FLNA) is a cytoplasmic protein that modulates cytoskeletal dynamics by crosslinking actin filaments to form branching actin networks. It also participates in the anchoring of actin filament bundles to membrane proteins, and thus plays a central role in cell mobility and structural integrity (Feng and Walsh 2004, Nakamura, Pudas et al. 2006, Yue, Huhn et al. 2013). To confirm that FLNA and NBEAL2 interact, 3XFlag-EGFP-NBEAL2 construct was transfected into HEK293 cells and NBEAL2 was immunoprecipitated with a GFP antibody. Immunoblot analysis confirmed that FLNA co-immunoprecipitated with NBEAL2, thus NBEAL2 was able to pull down endogenous FLNA (Fig. 4.4A), suggesting a positive interaction between NBEAL2 and FLNA. In light of this result, Dr. Hervé Falet (Blood Research Institute, Milwaukee, USA) has kindly provided us with FLNA consensus binding sequence that is found in FLNA interacting proteins. As shown in Fig. 4.4B, the FLNA binding consensus OXSXOXOXOXP (O = neutral amino acid residues, X = any amino acid residues, S = serine, P = proline) is found in 2740-2750aa of NBEAL2, with the exception of the first amino acid in the consensus being a basic glutamine instead of a neutral amino acid.

Figure 4.4. NBEAL2 interacts with FLNA. (A) 3XFlag-EGFP-NBEAL2 was transfected into HEK293 cells, NBEAL2 was immunoprecipitated with a GFP antibody and the eluate was analyzed with immunoblot. The result indicates an interaction between NBEAL2 and endogenous FLNA. (B) Recognition motif analysis revealed that 2740-2750aa of NBEAL2 may bind to FLNA. (Green = neutral amino acid residues, red = conserved amino acid residues).

14-3-3 proteins are a family of essential and ubiquitous proteins that regulate critical cellular processes including but not limited to apoptosis, signal transduction, cell cycling, cell development and transcription. 14-3-3 proteins are thought to bind to key proteins in various processes, altering their conformation and thereby regulating their function. They can also act as scaffold proteins to regulate cellular processes (Fu, Subramanian et al. 2000, Morrison 2009). Common recognition motifs for 14-3-3 proteins are phosphorylated sites depicted by RSXpS/TXP and RXXXpS/TXP, where X can be any amino acid residue and pS/T is either phosphorylated serine or threonine (Johnson, Crowther et al. 2010). These sites are

98 commonly found in WD40 domains where NBEAL2 contains five WD40 domains. In humans, seven isoforms of 14-3-3 were identified: β, γ, σ, ζ, ϵ, η, and τ. While deletion in one of the two isoforms of 14-3-3 proteins (BMH1 and BMH2) is tolerable in yeast, a double knockout is lethal. This suggests potential redundancy and overlapping functions between the 14-3-3 isoforms (Roberts, Mosch et al. 1997). Since all 14-3-3 proteins were found and are of the highest hit results obtained from the NBEAL2 affinity purification mass spectrometry, we proceeded with the verification of this interaction.

To detect interaction between 14-3-3 proteins and NBEAL2, primary human MKs were cultured, lysed and 14-3-3 proteins (β, γ, σ, ζ) were immunoprecipitated from the lysate with an antibody. Eluates were then analyzed with immunoblots. Results indicate that 14-3-3 proteins and NBEAL2 interact at the endogenous protein level where native levels of 14-3-3 proteins pulled down endogenous NBEAL2 (Fig. 4.5A). Primary human MKs were then examined with immunofluorescence confocal microscopy for subcellular co-localization of NBEAL2 and 14-3-3 proteins. 14-3-3 proteins are highly abundant, and as expected they appeared to be distributed uniformly throughout cells, while NBEAL2 showed a more punctate staining pattern (Fig. 4.5B). As a result, co-localization between 14-3-3 proteins and NBEAL2 is not apparent. Despite the lack of co-localization between NBEAL2 and 14-3-3 proteins, the interaction between NBEAL2 and 14-3-3 proteins is genuine. However, since 14-3-3 proteins are involved in a wide variety of processes, we continued to examine additional interacting partners that may offer more concrete insights into NBEAL2’s function in α-granule biogenesis.

Figure 4.5. NBEAL2 interacts with endogenous 14-3-3 proteins. (A) Primary human MKs were cultured and lysed, 14-3-3 proteins were immunoprecipitated and immunoblot analysis was performed with probing for NBEAL2. The results indicate endogenous 14-3-3 proteins interact with NBEAL2. (B) Immunofluorescence confocal microscopy analysis of the subcellular distribution of 14-3-3 proteins and NBEAL2 in human primary MKs. 14-3-3 appears to be distributed uniformly throughout cell while NBEAL2 shows a more distinct localization pattern. (Z-slice, scale bar = 5 µm).

4.3.4 NBEAL2 and SEC22B co-localize in primary human megakaryocytes

SEC22B is a SNARE protein involved in the transport of vesicles between the ER and Golgi (Liu, Flanagan et al. 2004). Recent advances revealed that SEC22B plays key roles in the maturation of phagosomes through endoplasmic reticulum-Golgi intermediate compartments (ERGIC)-phagosome vesicular transport and plasma membrane (PM) expansion through the formation ER-PM contact sites (Cebrian, Visentin et al. 2011, Petkovic, Jemaiel et al. 2014).

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Two methods were utilized to verify the interaction between NBEAL2 and SEC22B. First, HA- NBEAL2 construct was transfected into HEK293 cells, endogenous SEC22B was immunoprecipitated from the lysate, and immunoblot analysis was performed to detect HA- NBEAL2. Results indicate that transfected HA-NBEAL2 is able to pull down native SEC22B (Fig. 4.6A). Second, to ensure that the NBEAL2-SEC22B interaction in MKs was not a consequence of NBEAL2 over-expression, native protein immunoprecipitation was performed using primary human MK lysates. Native SEC22B was immunoprecipitated, and immunoblot analysis showed that native NBEAL2 co-immunoprecipitated (Fig. 4.6B). These co- immunoprecipitation experiments confirm that NBEAL2 interacts with SEC22B within primary MKs at endogenous protein expression levels.

To further examine the relationship between NBEAL2 and SEC22B, confocal immunofluorescence microscopy was performed using primary human MKs to visualize the intracellular localization of NBEAL2 and SEC22B. This analysis revealed a strong colocalization between NBEAL2 and SEC22B, with a Pearson’s correlation coefficient value of 0.681േ 0.63. Substantial overlap is also seen in the co-localization channel (Fig. 4.6C). Taken together, these results substantiate an interaction between NBEAL2 and SEC22B.

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Figure 4.6. NBEAL2 interacts with SEC22B. (A) HEK293 cells were transfected with HA-NBEAL2 and native SEC22B was immunoprecipitated with an antibody. Immunoblot was then probed with an NBEAL2 antibody. The analysis revealed an interaction between transfected NBEAL2 and native SEC22B. (B) Primary human MKs were lysed and SEC22B was immunoprecipitated with an antibody. Immunoblot analysis indicates that NBEAL2 interacts with SEC22B at the endogenous level in human primary MKs. (C) Confocal microscopy images of the subcellular localization of NBEAL2 and SEC22B in human primary MKs. Analysis revealed strong co-localization between NBEAL2 and SEC22B with a Pearson’s coefficient of 0.681േ 0.063 (n=3 independent experiments with 20-30 cells for each experiment; Z-slice, scale bar = 10 µm).

4.3.5 Delineation of binding region within NBEAL2 to SEC22B

With the NBEAL2-SEC22B interaction confirmed, we proceeded to investigate the details of binding. Specifically, we asked how NBEAL2 binds to SEC22B. The aim was to identify the critical regions within NBEAL2 that are responsible for interacting with SEC22B. Once these regions are identified, we can then begin to uncover the basis of the NBEAL2-SEC22B interaction and determine how this may relate to NBEAL2’s function during α-granule biogenesis. For instance, if the binding region of NBEAL2 to SEC22B overlaps with known missense mutations that cause GPS as shown in Fig. 4.7, we can then test whether these missense mutations impact

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NBEAL2-SEC22B interaction. Furthermore, we can then ask whether the GPS associated with these mutations is a result of the loss of interaction between NBEAL2 and SEC22B. While NBEAL2 is a 302 kDa protein with numerous domains, SEC22B is a small 25 kDa v-SNARE containing only a few domains: a longin domain, previously described as a negative regulator of membrane trafficking events, a SNARE domain for vesicle fusion and a transmembrane domain (TM) (Petkovic, Jemaiel et al. 2014). Although we are also interested in the binding region within SEC22B to NBEAL2, we focused our initial efforts on uncovering the regions within NBEAL2 that binds to SEC22B. Domain diagrams of NBEAL2 and SEC22B are shown in Fig 4.7.

Figure 4.7. Domain diagrams of NBEAL2 and SEC22B. NBEAL2 is a 302 kDa protein containing two armadillo-like domains (ARM) a concanavalin A-like lectin domain (ConA), a pleckstrin homology domain (PH), a BEACH domain and 5 WD40 domains. Missense mutations that have been reported to cause GPS are labelled in red. SEC22B is a 25 kDa protein containing a longin domain, SNARE domain and a transmembrane domain (TM).

We began by establishing a system to assess NBEAL2-SEC22B interactions. HA- NBEAL2 and Flag-SEC22B constructs were generated and transfected into HEK293 cells. Lysates were incubated with either a HA or Flag antibody, the eluates were processed for immunoblotting then probed with the corresponding antibodies to detect immunoprecipitation. It is evident based on the results that Flag-SEC22B co- immunoprecipitated with HA-NBEAL2 (Fig. 4.8A) and as expected, HA-NBEAL2 was also detected when Flag-SEC22B was immunoprecipitated (Fig. 4.8B). These results indicate that SEC22B and NBEAL2 interactions can be detected regardless of the direction of immunoprecipitation.

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Figure 4.8. Confirmation of NBEAL2-SEC22B interaction. HEK293 cells were transfected with HA- NBEAL2 and Flag-SEC22B. (A) HA-NBEAL2 was immunoprecipitated and immunoblot was probed with Flag antibody, followed by HA antibody to confirm pulldown efficiency. HA-NBEAL2 was capable of pulling down Flag-SEC22B. (B) Reversely, Flag-SEC22B was immunoprecipitated and the immunoblot was probed with HA antibody, followed by Flag antibody. Taken together the results indicate that NBEAL2 and SEC22B interact.

After establishing a working screening system, we began to delineate the binding region within NBEAL2 to SEC22B. Therefore, constructs that represent different portions of NBEAL2 were generated. As shown in Fig. 4.9, these expression constructs were engineered with a N-terminal HA-tag and are denoted C1 (1-550aa), C2 (530-900aa), C3 (877-1403aa), C4 (1380-1903aa), C5 (1877-2400aa) and C6 (2384-2754aa). These NBEAL2 constructs were transfected into HEK293 cells along with a Flag-SEC22B construct. Transfected cells were lysed, incubated with Flag M2 beads, then processed for immunoblotting to determine the regions of NBEAL2 that binds to SEC22B. Assessing the immunoblots, it is revealed that C1, C4 and C6 can immunopreciptate SEC22B (Fig. 4.9).

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Figure 4.9. Mapping the binding regions of NBEAL2 to SEC22B. HEK293 cells were co-transfected with Flag-SEC22B and one of the C1-C6 constructs where each represents a distinct portion of the full- length (FL) NBEAL2. HA-tagged proteins were immunoprecipitated and the immunoblot was probed with Flag antibody to detect binding followed by HA antibody to confirm the immunoprecipitation. The results suggest that SEC22B may interact with C1, C4 and C6, which represent 1-550aa, 1380-1903aa and 2384-2754aa of NBEAL2 respectively. * indicates IgG.

4.3.6 NBEAL2 contains an inhibitory domain that suppresses NBEAL2-SEC22B interaction

To determine whether a more complete NBEAL2 fragment influences the NBEAL2-SEC22B interaction, larger NBEAL2 fragment that encompass a higher degree of domain complexity, C7 (1-1145aa), C8 (1798-2754aa) and C9 (1380-2754aa), were tested. Based on the results, C7 does not interact with SEC22B (Fig. 4.10) despite C1, which is a region of NBEAL2 contained by C7, does (Fig. 4.9). C8 binds to SEC22B as expected since both C4 and C6, regions of NBEAL2 encompassed by C8, were positive in binding to SEC22B. Surprisingly, despite C8 binding to SEC22B, C9, a larger construct of C8 that is 418aa longer than C8 from the N-terminal, is incapable of binding to SEC22B (Fig. 4.10). This result suggests that NBEAL2 may contain an inhibitory region from 1380-1798aa and the expression of this region suppresses the NBEAL2-SEC22B interaction. An alternative explanation is that the inclusion of 1380-1798aa to C8 may have resulted in the formation of a WD40 propeller structure. In this scenario, the one WD40 domain from 1380-1798aa of NBEAL2 interacts with the four WD40 domains located at the C-terminal of C8, and as such, may have prevented SEC22B from being able to access NBEAL2. The sum of effects from the inhibitory domain and the formation of WD40 propeller structure may have resulted in a stronger suppression of NBEAL2-SEC22B interaction than that seen in C4, where C4 can still bind to SEC22B despite also harboring an inhibitory domain.

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To further examine this phenomenon, a construct expressing myc-tagged 1380-1798aa of NBEAL2, denoted by C10, was generated and co-transfected into HEK293 cells with FL-HA- NBEAL2 and Flag-SEC22B constructs for which an immunoprecipitation was then performed with a HA antibody. The results indicate that the C10, when over-expressed, inhibits the NBEAL2-SEC22B interaction (Fig. 4.11, bottom right panel). Based on this finding, the expression of C10 may be useful in introducing acute abrogation of NBEAL2-SEC22B interaction in cells. However, it should be noted that the inhibitory function of C10 was inconsistent in repeated trials of co-immunporecipitation experiments and therefore will require further testing prior to application.

We next wanted to delineate a more precise binding region of NBEAL2 with SEC22B. We showed that both C4 (1380-1903) and C6 (2384-2754aa) can interact with SEC22B (Fig. 4.9). Since no reports to date have indicated critical amino acid residues in the C6 region of NBEAL2, we focused our efforts on C4, which contains two sites that are known to cause GPS, E1833K and R1839C (Fig. 4.7). We excluded the 1380-1798aa from C4 (1380-1903aa) as this region was shown to be inhibitory to NBEAL2-SEC22B interaction. It is interesting to note that C4 despite containing the inhibitory domain is still capable of interacting with SEC22B, suggesting a potentially strong binding site contained within this fragment. C11 (1798-1903aa), a NBEAL2 fragment derived from C4 excluding the inhibitory region, was examined for interaction with SEC22B. The results indicate a strong interaction between C11 and SEC22B (Fig. 4.11, bottom center panel) while C10 does not interact with SEC22B (Fig. 4.11, bottom left panel), suggesting that the binding site of NBEAL2 to SEC22B is located within 1798-1903aa of NBEAL2. Similar to the co-expression of C10 disrupting the interaction between FL-NBEAL2 and SEC22B, introduction of over-expressed C10 is capable of suppressing the interaction between C11 and SEC22B (Fig. 4.11, bottom center panel).

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Figure 4.10. NBEAL2 contains inhibitory elements that prevent its interaction with SEC22B. HEK293 cells were transfected with Flag-SEC22B and one of the C7-C9 constructs where each represents a distinct portion of the full-length (FL) NBEAL2. C9 encompasses all five of the WD40 domains while C8 lacks one of the WD40 domain at its N-terminal. HA-tagged NBEAL2 fragments were immunoprecipitated and the immunoblot was probed with Flag antibody to detect binding to SEC22B. Immunoblots were then probed with HA antibody to assess pulldown efficiency. The results indicate that C8 binds to SEC22B while C9, which contains additional sequences at its N-terminal relative to C8, fails to interact with SEC22B.

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Figure 4.11. Defining the binding regions of NBEAL2 and SEC22B. HEK293 cells were transfected with Flag-SEC22B and NBEAL2 constructs. NBEAL2 constructs were immunoprecipitated with HA antibody and the immunoblot was probed with Flag antibody to detect binding with SEC22B and HA antibody to confirm pulldown efficiency. Results revealed that C10 (1380-1797aa of NBEAL2) does not interact with SEC22B while C11 (1798-1903aa of NBEAL2) interacts with SEC22B (lower left and lower center panels). Co-expression of C10 (1380-1797aa of NBEAL2) disrupts NBEAL2-SEC22B and C11-SEC22B interactions (lower center and lower right panels). Myc-C10 expression was not shown as the protein band overlaps directly with IgG bands and was indistinguishable.

4.3.7 NBEAL2 protein level is dependent on the presence of SEC22B

To explore the functional significance of SEC22B binding to NBEAL2’s in α-granule biogenesis, we utilized immortalized megakaryocyte progenitor (imMKCL) cells. ImMKCL is a megakaryocytic cell line established from human pluripotent cells obtained from Dr. Koji Eto in Japan (Nakamura, Takayama et al. 2014). We have validated that imMKCL cells were capable of forming α-granules as analyzed by TEM. We then proceeded to utilize the CRISPR/Cas9 system to knockout NBEAL2 and SEC22B in these imMKCL cells to determine if the knockout causes GPS. To accomplish this, a lentivirus protocol was developed to introduce CRISPR/Cas9 into these cells as conventional transfection techniques have proved highly ineffective. This method was effective in the generation of NBEAL2 and SEC22B KO imMKCL cell lines. Interestingly, the levels of NBEAL2 were significantly reduced when SEC22B was knocked out, indicating that the level of NBEAL2 is dependent on the presence of SEC22B (Fig. 4.12A). VWF level is lowered in both of the knockout cell types. Densitometry analysis revealed that NBEAL2 level is significantly lowered when SEC22B is

108 knocked out (Fig. 4.12B) whereas knockout of NBEAL2 has no significant effect on the level of SEC22B (Fig. 4.12C).

Figure 4.12. Expression of NBEAL2 is dependent on SEC22B. ImMKCL cells were transduced with lentivirus packaged with constructs designed to knockout NBEAL2 or SEC22B via CRISPR. Positive cells were expanded following puromycin selection and analyzed via immunoblot. (A) Immunoblot confirms the knockout of NBEAL2 and SEC22B in the respective imMKCL cell populations. VWF

109 expression was lower in both of the knockout cell lines. Interestingly, NBEAL2 expression is reduced when SEC22B was absent, suggesting that the expression of NBEAL2 is reliant on SEC22B expression. (B) Densitometric analysis indicates that the level of NBEAL2 is significantly reduced when SEC22B is knocked out while (C) NBEAL2 knockout does not affect SEC22B expression. Data representative of 3 independent experiments; error bars = standard deviation, * = P<0.05 via student’s t-test, NS = p>0.05.

4.3.8 NBEAL2 and SEC22B are both required for α-granule biogenesis

To determine the significance of SEC22B for the formation of α-granules, SEC22B KO imMKCL cells were examined with TEM for the presence of α-granules. It is anticipated that if SEC22B is not required for α-granule biogenesis, the loss of SEC22B would have no effects on the α-granules in the cell. Analysis was performed in parallel with WT and NBEAL2 KO imMKCL cells. TEM of these cell lines revealed that both NBEAL2 and SEC22B KO imMKCL cell lines exhibited α-granule deficiencies, a characteristic of GPS (Fig. 4.13). These results suggest that SEC22B is essential for α-granule biogenesis where it likely involves an interacion with NBEAL2. SEC22 loss leads to a GPS-like phenotypes in imMKCL cells similar to that seen in NBEAL2 KO imMKCL cells.

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Figure 4.13. SEC22B is required for α-granule formation in ImMKCL cells. The ultrastructures of WT, NBEAL2 KO and SEC22B KO imMKCL cells were visualized with transmission electron microscopy (TEM). Numerous electron dense α-granules were observed in WT imMKCL cells while these structures were absent when either NBEAL2 or SEC22B was absent. These results indicate that SEC22B is required for α-granule formation similar to NBEAL2.

4.3.9 NBEAL2-SEC22B interaction is essential for α-granule formation

While we have identified that knocking out either NBEAL2 or SEC22B leads to GPS-like phenotypes in imMKCL cells and thereby confirming SEC22B as a critical protein in α-granule biogenesis, it remains unclear whether the phenotypes observed were linked specifically to the disruption of the interaction between NBEAL2 and SEC22B. To further explore the properties of NBEAL2-SEC22B interaction, we screened for the GPS-causing NBEAL2 mutations that are located within the amino acid residues, 1798-1903, of NBEAL2. This is the region of NBEAL2 that was identified to bind to SEC22B, as described in Fig. 4.11. Of the NBEAL2 missense mutations that cause GPS, E1833K and R1839C are located within the 1798-1903aa binding region. To determine whether these mutations can potentially interfere

111 with NBEAL2-SEC22B interaction, NBEAL2 constructs harboring these mutations were generated and co-immunoprecipitation experiments were performed in HEK293 cells to test for their association with SEC22B. The results shown in Fig. 4.14A reveal that both E1833K and R1839C abrogated the NBEAL2-SEC22B interaction. A control co-immunoprecipitation was performed with VAP-A, a NBEAL2 interacting protein that was obtained from the affinity purification mass spectrometry as described previously in Table 4.3, where the results suggested that the E1833K and R1839C mutations are likely specific to NBEAL2-SEC22B interaction (Fig. 4.14B). These findings support the importance of the NBEAL2-SEC22B interaction for α-granule biogenesis and may account for the cause of GPS seen in patients with these NBEAL2 mutations.

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Figure 4.14. NBEAL2 E1833K and R1839C mutations abrogate NBEAL2-SEC22B interaction. E1833K and R1839C missense mutations in NBEAL2 are known to cause GPS and are both located in the region where NBEAL2 interacts with SEC22B. (A) HA-NBEAL2 E1833K or R1839C mutation constructs were transfected into HEK293 cells alongside with Flag-SEC22B. NBEAL2 was immunoprecipitated and analyzed with immunoblot for interaction with SEC22B. Results indicate that both E1833K and R1839C mutations in NBEAL2 abrogate NBEAL2-SEC22B interaction. (B) E1833K and R1839C missense mutations did not prevent NBEAL2-VAPA interaction, suggesting that these mutations are specific to NBEAL2-SEC22B binding.

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4.4 Discussion 4.4.1 Binding partners provide insights into NBEAL2 function

Through affinity purification – mass spectrometry (AP-MS), potential interactors of NBEAL2 were identified. Among the candidates, 14-3-3, FLNA and SEC22B were confirmed to interact with NBEAL2. It was expected that several NBEAL2 binding partners were going to be identified through this assay since BDCP were proposed to function as scaffold proteins (Cullinane, Schaffer et al. 2013). Furthermore NBEAL2 is a large protein that contains multiple homology domains that are known to function in protein-protein interactions, such as the ARM and WD40 domains (Hatzfeld 1999, Xu and Min 2011). These features allow NBEAL2 to potentially interact with a diverse range of proteins via the different interacting domains, effectively making it a scaffold protein. As I have shown here, NBEAL2 appears to interact with a wide-range of proteins, each with different known functions. This suggests that NBEAL2 function in MKs may not be limited to α-granule biogenesis. This view is supported by the various MK and platelet defects seen in GPS patients and Nbeal2-/- mouse models in addition to the α-granule deficiency. These include impairments in megakaryopoiesis, thrombopoiesis and DMS development (Gunay-Aygun, Falik-Zaccai et al. 2011, Deppermann, Cherpokova et al. 2013, Guerrero, Bennett et al. 2014). Therfore, the identification of these proteins will provide further insights into NBEAL2 function once more precise roles of the NBEAL2-interacting proteins during MK development are determined.

14-3-3 is a family of scaffold proteins implicated in diverse cellular processes including apoptosis, signal transduction, cell cycling and development (Fu, Subramanian et al. 2000, Morrison 2009). Without prior knowledge, it is difficult to pinpoint the implication of NBEAL2- 14-3-3 interaction in MKs due to the diverse functions of 14-3-3. However, among the diverse functions of 14-3-3, It is well recognized that 14-3-3 mediated cell signaling events are critical to cell survival, development and growth (Dong, Kang et al. 2007). As Nbeal2-/- MKs displayed delayed development and abnormal DMS formation, it is possible that the interaction between 14-3-3 and NBEAL2 may mediate these cellular process (Kahr, Lo et al. 2013, Guerrero, Bennett et al. 2014). Further, platelet activation has been shown to be intricately correlated with the activity of 14-3-3 (Chen, Ruggeri et al. 2018), suggesting a potential role of NBEAL2 in modulating platelet activity via 14-3-3. FLNA is an actin-associated protein involved in the modelling of cellular cytoskeleton. Interestingly, macrothrombocytopenia and abnormal DMS formation in MKs are phenotypes shared by both the Nbeal2-/- mice and FLNA-/- mice (Kahr, Lo et al. 2013, Begonja, Pluthero et al. 2015). This suggests a potential significance of

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NBEAL2-FLNA interaction in regulating thrombopoiesis and DMS development. It is possible that the failure of NBEAL2 to interact with 14-3-3 and FLNA leads to some of the defects seen in Nbeal2-/- mice. As the clinical manifestation of GPS varies greatly among patients (Nurden and Nurden 2007, Gunay-Aygun, Zivony-Elboum et al. 2010), it is conceivable that different NBEAL2 missense mutations result in different protein interaction failures, and thereby provides the heterogeneity of phenotypes seen in GPS patients.

SEC22B is an ER protein that was first implicated to function as a v-SNARE to regulate both the anterograde and retrograde transport of vesicles between the ER and Golgi (Zhang, Wong et al. 1999, Mossessova, Bickford et al. 2003, Mancias and Goldberg 2007). Since then, additional roles of SEC22B have been uncovered. Most notably, it has been shown that SEC22B is vital for phagosome maturation in dendritic cells. In this process, SEC22B regulates the transport of phagosomal contents from the ERGIC to the phagosomes, facilitating their maturation. Depletion of SEC22B inhibits phagosome function and results in the abnormal presentation of antigens by dendritic cells (Cebrian, Visentin et al. 2011). By analogy, it is possible that SEC22B may function in the vesicular transport of α-granule cargoes. In addition, SEC22B is also implicated in the expansion of the PM. Through the formation of MCS with syntaxin-1 on the PM, SEC22B facilitate lipid transfer from the ER to the PM to promote cell expansion and growth (Petkovic, Jemaiel et al. 2014). With ER- endosome MCS being implicated in the maturation of endosomes, this suggest a potential role of SEC22B in regulating α-granule maturation (Friedman, Dibenedetto et al. 2013). It is also possible that formation of MCS between the ER and α-granules provides structural support to α-granules and thereby promoting their stability. While it is unclear which pathway is implicated by NBEAL2 and SEC22B during α-granule biogenesis in MKs, these studies nonetheless suggest that NBEAL2 and SEC22B may act in tandem to regulate α-granule maturation.

It is important to note that this interactome study was performed from stable DAMI cell lysates. While DAMI cells are megakaryocytic in nature, these cells do not appear to produce proper α-granules when examined via TEM. This suggests the possibility that some key NBEAL2 interactions that are relevant for α-granule biogenesis may be missing from this interactome study but may be present in native MKs. This is likely the case for P-selectin. In our NBEAL2 interactome study in DAMI cells, P-selectin was not identified as one of the interacting partners of NBEAL2. However, endogenous immunoprecipitation of NBEAL2 from human

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MKs and platelets indicate an authentic interaction between NBEAL2 and P-selectin (Fig. 3.10 and Fig. 3.11). Despite this caveat, some NBEAL2 interactions in DAMI cells are likely genuine since it is a megakaryocytic cell line that has some features of MKs such as the production of the α-granule cargo protein VWF. Thus, identification of NBEAL2 interacting proteins in DAMI cells has nonetheless yielded valuable insights into the functions of NBEAL2.

A similar interactome study of NBEAL2 was performed in HEK293 cells. In this study, DOCK, VAC14 and SEC16A were identified as binding partners of NBEAL2 (Mayer, Jasztal et al. 2018). DOCK7 is a cytoplasmic guanine exchange factor (GEF) involved in the binding and activation of RAC GTPases such as RAC1 (Watabe-Uchida, John et al. 2006, Sobczak, Chumak et al. 2016, Nakamuta, Yang et al. 2017), a positive regulator of platelet granule secretion. (Akbar, Kim et al. 2007, Flevaris, Li et al. 2009) Thus DOCK7-NBEAL2 interactions may negatively regulate α-granule secretion in MKs and/or platelets. VAC14 is a regulator of phosphatidylinositol 3,5-bisphosphate (PI(3,5)P2) that localizes to a wide range of endocytic organelles (Zhang, McCartney et al. 2012), possibly including α-granules. Loss of VAC14 leads to vacuolated neurons, (Zhang, Zolov et al. 2007) thus interactions with NBEAL2 may contribute to the maturation and/or maintenance of α-granules. SEC16A is a membrane protein located at ER exit sites that is implicated in the anterograde trafficking of COPII vesicles from ER to Golgi (Budnik and Stephens 2009). ER-endosome membrane contact sites are important in endosome maturation (Friedman, Dibenedetto et al. 2013), and it is possible that SEC16A-NBEAL2 interactions facilitate membrane transfers from ER to nascent α-granules. However, it is also important to note that HEK293 cells are not representative of MKs. Therefore, NBEAL2 interactions in HEK293 cells may be different from that in MKs or megakaryocytic cell lines (DAMI). This may be one of the underlying factors accounting for the differences observed in our NBEAL2 interactome study as we did not identify DOCK7, VAC14 or SEC16A.

4.4.2 Dynamic interaction between NBEAL2 and SEC22B

To understand the nature of the interaction between NBEAL2 and SEC22B, we delineated the binding region within NBEAL2 to SEC22B. From these assays, it was determined that C11, constituting 1798-1903aa of NBEAL2, interacts with SEC22B. Outside of C11, potential interaction with SEC22B were also observed for C1 (1-550aa of NBEAL2) and C6 (2384- 2754aa). However, C1-SEC22B interaction was not pursued as the extended version of C1,

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C7 (1-1145aa of NBEAL2), did not bind to SEC22B. This suggests a scenario of a more complicated interaction at this region. The interaction between C6 and SEC22B was retained as there are currently no GPS-causing mutations being reported in this region of NBEAL2. While it cannot be excluded that the binding of SEC22B to this region of NBEAL2 is important to the functions NBEAL2, it is interesting that C11 contains two reported GPS-causing mutations, E1833K and R1839C, which abrogate binding to SEC22B. This suggests that this region of NBEAL2 contains critical amino acids that may be related to its interaction with SEC22B, and hence function for α-granule formation.

From our study, we also propose that the interaction between NBEAL2 and SEC22B may be regulated by NBEAL2 itself. The basis of this proposal stems from the observation that while C8 (1798-2754aa of NBEAL2) can bind to SEC22B effectively, an extended version of C8, C9 (1380-2754aa of NBEAL2), fails to bind to SEC22B. This suggest that a potential inhibitory domain exists within the region of 1380-1798aa of NBEAL2. However no firm conclusions can yet be made as there were inconsitent results with some repeat experiments. If 1380-1798aa of NBEAL2 does indeed contain an inhibitory element, one needs to consider how C4 (1380- 1903aa of NBEAL2), which contains the putative inhibitory region, can still bind to SEC22B while C9 (1380-2754aa of NBEAL2), an extended version of C4 which also contains the putative inhibitory region, fails to bind to SEC22B. Comparing C4 and C9, C4 contains one WD40 domain (1515-1555aa of NBEAL2) while C9 contains five WD40 domains, one from 1515-1555aa of NBEAL2 and four from 2457-2724aa of NBEAL2. As WD40 β-propeller is generally formed from four to sixteen WD40 blade repeats (Smith, Gaitatzes et al. 1999, Li and Roberts 2001, Xu and Min 2011), C9 is capable of forming WD40 β-propeller while C4 cannot. The formation of WD40 β-propeller between the one WD40 repeat at the N-terminal (1515-1555aa of NBEAL2) and four WD40 repeats at the C-terminal (2457-2724aa of NBEAL2) of C9 may have prevented SEC22B from accessing the binding region that is located in between the WD40 repeats (1798-1903aa of NBEAL2). Thus, as opposed to C4 which only contains a putative inhibitory domain, the combination of the putative inhibitory domain and the formation of WD40 propeller in C9 may produce a more complete regulatory structure to modulate the NBEAL2-SEC22B interaction.

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4.4.3 SEC22B affects NBEAL2 protein levels

Analysis of imMKCL cells has revealed that the NBEAL2 level is decreased when SEC22B is absent, suggesting that either the expression levels or the stability of NBEAL2 is dependent on the presence of SEC22B. This phenomenon highlights and reinforces the importance of the NBEAL2-SEC22B interaction for NBEAL2 function during α-granule biogenesis. Conversely, knocking-out NBEAL2 does not affect the level of SEC22B in imMKCL cells. This is reasonable, since SEC22B is a protein that is expressed at a much higher level than NBEAL2 in platelets (Simon, Edelstein et al. 2014), suggesting that SEC22B likely has multiple functional roles in MKs relative to NBEAL2. This is consistent with the role of SEC22B as an ER protein that is likely involved in multiple cellular processes independent of NBEAL2. Hence, these data indicate a dependence of NBEAL2 on SEC22B but not SEC22B on NBEAL2.

4.4.4 NBEAL2 and SEC22B are critical for α-granule biogenesis

To determine whether SEC22B is vital to the function of NBEAL2, a follow-up study examined the significance of SEC22B for α-granule biogenesis. TEM analysis of SEC22B-KO imMKCL cells revealed that similar to NBEAL2-KO imMKCL cells, SEC22B-KO imMKCL cells recapitulated GPS as they lack α-granules. This indicated that knocking out SEC22B results in the same phenotype as knocking out NBEAL2, suggesting that both proteins are required for α-granule biogenesis. Since NBEAL2 and SEC22B interact, they may act in tandem during α-granule biogenesis. This result highlights the significance of SEC22B and the potential importance of the NBEAL2-SEC22B interaction for α-granule biogenesis. A caveat to this study is the potential off-target effects of CRISPR that may have been a confounding variable. This phenomenon in itself may be interesting if true, since few genes have been implicated in α-granule biogenesis. Ideally, the reintroduction of SEC22B to rescue the induced α-granule defects in SEC22B-KO imMKCL cells would serve as a useful control. Regardless, the evidence that SEC22B affects NBEAL2 protein levels, suggests that SEC22B is a relevant molecule required for α-granule biogenesis.

To understand the importance of the NBEAL2-SEC22B interaction for α-granule biogenesis further, we analyzed missense mutations in NBEAL2 that have previously been reported to cause GPS. We showed that C11 (1798-1903aa of NBEAL2) interacts with SEC22B and contains two GPS-causing missense mutations (E1833K and R1839C). This allowed us to

118 determine the significance of the NBEAL2-SEC22B interaction for α-granule biogenesis. Interestingly, NBEAL2 containing either E1833K or R1839C mutations failed to bind with SEC22B, confirming the importance of the NBEAL2-SEC22B interaction for α-granule biogenesis. To further interrogate the requirement for the NBEAL2-SEC22B interaction for α- granule biogenesis, a follow-up approach could be to try to rescue the α-granule defect in NBEAL2-KO imMKCL cells using SEC22B-binding deficient NBEAL2. The prediction would be that if the NBEAL2-SEC22B interaction is vital to α-granule biogenesis, the introduction of SEC22B-binding deficient NBEAL2 would not rescue the α-granule defects. This approach would provide further insights into the significance of the NBEAL2-SEC22B interaction in α- granule biogenesis.

While additional experimentation may reinforce the hypothesis that the NBEAL2-SEC22B interaction is required for α-granule biogenesis, our data have already provided significant insights into this process. Currently, two potential models exist for the action of SEC22B. First, SEC22B may associate with NBEAL2 to mediate the trafficking of α-granule cargoes, thereby regulating the maturation of α-granules. Alternatively, SEC22B and NBEAL2 may form MCS. Since SEC22B is an ER protein and NBEAL2 is associated with α-granules as previously shown by us, this suggests that the NBEAL2-SEC22B interaction could bring the ER and α- granules in close proximity. This interaction could enhance the stability and maturation of α- granules, potentially through lipid transfers from the ER to the α-granules. Taken together our study has provided important new insights into the biology of α-granule biogenesis. We have shown that SEC22B is an interactor of NBEAL2 and similar to NBEAL2, is also a critical protein in α-granule biogenesis, suggesting that SEC22B and NBEAL2 may function in tandem.

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Chapter 5

NBEAL2 is required for normal development of megakaryocytes

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5.1 Abstract

GPS is a bleeding disorder caused by mutations in NBEAL2 that is characterized by macrothrombocytopenia, bone marrow myelofibrosis and splenomegaly. The paucity of α- granules in GPS platelets associates NBEAL2 to α-granule formation. To investigate the role of NBEAL2 in MKs and platelets, we studied Nbeal2-/- mice as a model of GPS. Similar to human GPS, Nbeal2-/- mice display macrothrombocytopenia, splenolomegaly and platelet α- granule deficiency as characterized by the lack of α-granule cargo proteins. Interestingly, P- selectin, an α-granule membrane constituent, remains present, albeit at a reduced level in Nbeal2-/- platelets. Analysis of Nbeal2-/- platelets through flow cytometry, platelet aggregometry, bleeding assays and intravital imaging of thrombus formation revealed impaired platelet function. Using electron microscopy, Nbeal2-/- MKs showed abnormal structural organization and α-granule deficiency. Cultured Nbeal2-/- MKs displayed delayed development, decreased survival, decreased ploidy and decreased proplatelet production. Immunofluorescence analysis of Nbeal2-/- MKs revealed peripheralization of VWF, consistent with the increased presence of TSP1 in Nbeal2-/- MK culture medium. Together with the deficiency of α-granules in Nbeal2-/- MKs, these observations suggest that this is the MK developmental stage where α-granules are lost. Our study confirmed the vital importance of NBEAL2 for α-granule formation, megakaryopoiesis, thrombopoiesis and platelet function.

5.2 Introduction

GPS is an inherited bleeding disorder characterized by macrothrombocytopenia, bone marrow myelofibrosis, splenomegaly and paucity of platelet α-granules. While GPS has been linked to mutations in the gene encoding for NBEAL2 (Albers, Cvejic et al. 2011, Gunay- Aygun, Falik-Zaccai et al. 2011, Kahr, Hinckley et al. 2011), the functional role of NBEAL2 is unknown. As NBEAL2 is clearly essential for α-granule biogenesis, understanding the mechanistic details of NBEAL2 function, would provide valuable insights into the process of α-granule formation. Furthermore, potential alternative roles of NBEAL2 during MK and platelet development may be important. The study of α-granule biogenesis has been challenging due to the fact that only few diseases have been associated with α-granule deficiencies and the absence of a suitable model cell. In this study, we investigated the functions and roles of NBEAL2 in MKs and platelets by describing Nbeal2-/- mice, a model of human GPS (Kahr, Lo et al. 2013). The aim of this study was to gain insights into the pathway of α-granule biogenesis as well as uncovering the nature and function of NBEAL2.

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5.3 Results

Work in this chapter was published in Blood (Kahr, Lo et al. 2013), excluding 5.13, 5.14 and 5.15. TEM was performed by Hilary Christensen (Fig. 5.1D/E, 5.5 and 5.11D). Confocal microscopy of platelets was performed by Dr. Fred Pluthero (Fig. 5.3D). Platelet function analysis in Fig. 5.4 was performed by Dr. Fred Pluthero and members of Dr. Peter Gross’s lab (McMaster University).

5.3.1 Nbeal2-/- mice as a model of human GPS

Nbeal2-/- mice were generated to represent human GPS and to understand the role of NBEAL2 in MKs and platelets. Offspring were fertile, viable and did not present distinct abnormalities when compared with WT mice controls. Four month old mice were examined. Significant splenomegaly was observed in Nbeal2-/- mice (WT: 0.09േ0.02g; Nbeal2-/-: 0.16േ0.05g; P=0.0001, Mann-Whitney 2-tailed test; n=13 mice each condition), along with significantly increased MK counts within the spleen of Nbeal2-/- mice when examined with hematoxylin and eosin stain (WT spleen slice: 67േ19; Nbeal2-/- spleen slice: 330േ12; n=3 each condition; P<0.0001, 2-tailed t-test). Nbeal2-/- mice displayed characteristics of macrothrombytopenia where low platelet counts (WT: 848േ202*109/L; Nbeal2-/-: 519േ55* 109/L; n=3 mice per condition) and higher platelet volume (WT: 5.10േ0.85 fL; Nbeal2-/-: 6.69േ0.35 fL; n=3 mice per condition) were observed. Blood film analysis of WT and Nbeal2+/- mice platelets showed normal platelet staining (Fig. 5.1A and Fig. 5.1B respectively) whereas platelets from Nbeal2-/- mice were larger and pale in color (Fig. 5.1C). Platelet ultrastructure was examined with thin-section TEM and revealed the presence of α-granules in platelets from WT mice (Fig. 5.1D) but the absence of α-granules in Nbeal2-/- platelets (Fig. 5.1E). Evaluation of α-granules through morphometric analysis revealed an average of 0.25 α-granules per Nbeal2-/- platelet and 4.8 α-granules per WT platelet (n=25 platelets for WT and 100 platelets for Nbeal2-/-). Overall, these phenotypes are consistent with and reflect human GPS. This supports the use of Nbeal2-/- mice as a model to represent human GPS. Exceptions are the presence myelofibrosis and δ-granule counts. Analysis of Nbeal2-/- mouse bone marrow indicated a lack of myelofibrosis which is sometimes observed in GPS patients. While GPS platelets generally contain normal or increased δ-granule counts when examined with whole-mount electron microscopy, Nbeal2-/- platelets showed lower δ-granule counts in comparison to WT platelets (WT: 4.9 δ-granules per platelet; Nbeal2-/-: 2.6 δ-granules per platelet; n=50 platelets per condition).

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Figure 5.1. Blood film and EM ultrastructure abnormalities of Nbeal2-/- mouse platelets. Blood films were prepared from WT (A), Nbeal2+/- (B), and Nbeal2-/- (C) mice and stained with Wright-Giemsa stain prior to light microscopy. WT (A) and Nbeal2+/- (B) blood films show typical platelet (black arrowheads) size and morphology with discernible granulation against light cytoplasmic staining, and the Nbeal2-/- film (C) shows gray-appearing platelets with indistinct granulation and visible vacuoles. Thin section transmission electron micrographs of representative platelets are from WT mice (D) and Nbeal2-/- mice (E). Multiple α-granules (white arrowheads) were evident in WT platelets and absent in Nbeal2-/- platelets, which are larger on average. Magnification X40,000; scale bars=500 nm.

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5.3.2 Nbeal2-/- platelets contain reduced α-granule cargo and membrane proteins

Deficiency of platelet α-granules is a hallmark of GPS in humans. With confirmation from TEM that Nbeal2-/- mice platelets also lacked α-granules, we next want to assess the presence α-granule cargoes. Immunoblot analysis of Nbeal2-/- platelet lysates revealed the absence of α-granule cargo proteins including VWF and TSP1 (Fig. 5.2). In addition, knockout of NBEAL2 was confirmed using the same immunoblot. Densitometric analysis revealed that the expression of the α-granule membrane constituent, P-selectin, was lowered in Nbeal2-/- platelets relative to WT (48.4േ5.3% of WT). This suggests that α-granule membrane structures may still be present in these cells, however NBEAL2 may be required for maintaining normal P-selectin levels. This contrasts with ARC syndrome where the lack of VPS33B or VPS16B results in the complete loss of soluble α-granule cargo proteins and P- selectin in platelets. This difference between GPS and ARC syndrome suggests that VPS16B/33B may be required in an earlier phases of α-granule formation before NBEAL2.

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Figure 5.2. Granule cargo proteins, P-selectin and NBEAL2 in platelets and megakaryocytes. (A) Immunoblot analysis confirmed decreased α-granule borne proteins VWF, TSP1 and FGN, as well as granule membrane-associated P-selectin, in Nbeal2-/- mouse platelet lysates relative to WT (GAPDH used as protein loading control). (B) FGN is also reduced in Nbeal2-/- mouse platelet lysate relative to WT, and analysis of lysates from day 5 MKs cultured from bone marrow from both sources show absence of endogenous FGN, consistent with the necessity for α-granule FGN to be endocytosed by MKs. (C) Densitometric analysis indicate that the P-selectin level in Nbeal2-/- mice platelets is 48.4േ5.3% of WT levels (n = 3 independent experiments).

5.3.3 Structural and functional abnormalities of Nbeal2-/- megakaryocytes and platelets

With the establishment of Nbeal2-/- mice as a suitable model of human GPS, we next assessed whether MKs and platelets lacking NBEAL2 displayed any other structural and

125 functional abnormalities. Results from such studies could provide insights into the phenotypes seen in GPS and point to alternative functions of NBEAL2. Staining of Nbeal2-/- platelets showed a similar pattern of distribution of CD41 (integrin αIIb). While P-selectin distribution appeared more disorganized in Nbeal2-/- platelets, no striking differences were observed in comparison to WT (Fig 5.3A-B). Lack of NBEAL2 also does not prevent the mobilization of P- selectin towards the cell surface, indicating the cell is capable of α-granule secretion (Fig. 5.3C-D).

Figure 5.3. P-selectin in resting and thrombin-activated Nbeal2-/- platelets. High-resolution confocal laser immunofluorescence microscopy imaging of intracellular P-selectin membrane proteins in fixed resting platelets. Permeabilized cells were stained for α-tubulin (violet), P-selectin (red), and CD41/integrin aIIb (green) and imaged (final magnification = X150, Z stepping = 250 nm). Single- channel and merged midcell ZY/XY slices of representative individual platelets are shown (A-B). Resting platelets from WT (A) and Nbeal2-/- (B) mice have similar flat, discoid morphology with a well- defined circumferential tubulin ring cytoskeleton. Both contain P-selectin, which defines compact looping structures of the α-granule secretory matrix typical of WT platelets that generally appear to be 126 less orderly in Nbeal2-/- platelets. Thrombin activated WT (C) and Nbeal2-/- (D) platelets show characteristic activation-triggered changes in shape and surface mobilization of P-selectin.

To analyze platelet function, platelets were activated with various agonists and the level of activation was measured with flow cytometry by detecting aIIbβ3 activation with JON/A-PE or the presence of surface P-selectin. Measuring the levels of activated aIIbβ3, Nbeal2-/- platelets showed reduced response to both thrombin and ADP activation (Fig. 5.4A-B). Similar deficiency in activation was observed when Nbeal2-/- platelets were activated by thrombin and levels of externalized P-selectin were measured (Fig. 5.4C). Despite the levels of P-selectin in Nbeal2-/- platelets is 48.4േ5.3% relative to WT, maximum level of P-selectin exposure from Nbeal2-/- platelets was only one-third in comparison to WT, suggesting a fundamental impairment in P-selectin mobilization. Light emission aggregometry revealed Nbeal2-/- platelets are impaired in activation response when cells were activated with either collagen-related peptide (CRP) (Fig. 5.4D), thrombin (Fig.5.4E) or thromboxane-prostanoid receptor agonist U46619 (Fig. 5.4F). Consistently, impedance aggregometry using a Multiplate analyzer revealed Nbeal2-/- platelets have reduced activation capabilities when cells were exposed to collagen (Fig. 5.4G). These results are consistent and revealed a functional impairment in Nbeal2-/- platelets.

We next asked whether the functional impairments of Nbeal2-/- platelets observed in vitro were also reflected in vivo. Since platelets facilitate blood clotting, mice tails were bled to assess the rate of clotting. Tail transection assay revealed Nbeal2-/- mice displayed higher cumulative blood loss, suggesting deficiencies in clotting relative to WT (Fig. 5.4H). Further, thrombi formation was monitored with intravital microscopy following laser injury of cremaster muscle arterioles. Accumulation of platelets within thrombi were measured with fluorescent anti-GPIbβ antibody (X488) in terms of sum, intensity three minutes after injury, and maximal accumulation. Intravital microscopy revealed Nbeal2-/- platelets showed reduced functions in thrombi formation (Fig. 5.4I-J; WT: P=0.03; Nbeal2-/-: P=0.04). Interestingly, despite impairment of Nbeal2-/- platelets to accumulate at the thrombi, measurements of surface CD41 and P-selectin revealed that Nbeal2-/- platelets have similar rate of activation as WT platelets (Fig. 5.4K-L). However, since Nbeal2-/- platelets have a lower maximal P-selectin to GPIbβ ratio than WT platelets (Fig. 5.4M), these data suggest that Nbeal2-/- platelets have lower maximal P-selectin expression despite their rate of activation being similar to WT platelets.

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Figure 5.4. Impaired Nbeal2-/- mouse platelet function in vitro and in vivo. Measurements of activation of platelets from WT (solid symbols) and Nbeal2-/- mice (open symbols). (A-C) Washed platelets were assessed by flow cytometry for activation in response to varying concentrations of agonists (X-axis) by measuring antibody binding (JON/A) to activated aIIbb3 (A-B) or exposed P- selectin (C). Graphs show mean േ standard error of the mean (SEM) (n = 6 mice per group) for mean fluorescence intensity (MFI) after activation by thrombin (A,C) or ADP (B). (D-F) Optical aggregometry in physiological buffer of washed platelets exposed to varying concentrations of CRP (D), thrombin (E), or the thromboxane analog U46619 (F). The agonist concentration is plotted against the percentage of aggregation (see the “Methods” section). (G) Impedance aggregometry measurement of platelets in citrated blood exposed to collagen (7 mg/mL, n = 6 mice per group); horizontal lines represent mean േ SEM for area under the curve of Multiplate aggregation. (H) Nbeal2-/- mice showed greater cumulative blood loss in relation to WT in a tail transection bleeding assay; mean 6 SEM is shown for WT (solid squares) and Nbeal2-/- mice (open circles; n = 11 for each). (I-M) Results of intravital video microscopy monitoring of platelet accumulation and activation in thrombi formed in response to laser injury of cremaster muscle arterioles. (I-J) Sum and maximal platelet accumulation in thrombi formed in response to injury were determined using a fluorescent anti-mouse GPIbb antibody (X488); values shown are mean േ SEM for 170 thrombi in 12 WT mice and 155 thrombi in 12 Nbeal2-/- mice (*P < .05, unpaired t test). (K-L) The time to half-maximal activation ratio for CD41 was determined using an anti- mouse CD41 Fab fragment (K) and for P-selectin (L) using an anti-mouse P-selectin antibody. Values shown are mean േ SEM; for CD41, n = 119 thrombi in 6 WT mice and 93 thrombi in 6 Nbeal2-/- mice; for P-selectin, n = 69 thrombi in 6 WT mice and 60 thrombi in 6 Nbeal2-/- mice. (M) The time course of P-selectin activation after injury. Mean േ SEM for WT mice (solid squares) and Nbeal2-/- mice (open circles) are shown.

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Bone marrow MKs were examined to determine any impacts of Nbeal2 deletion. TEM analysis of bone marrow MKs from WT mice revealed the presence of typical α-granules and distinct platelet territories (Fig. 5.5A,C) whereas MKs from Nbeal2-/- bone marrow showed reduced numbers of typical α-granules and a lack of clear platelet territories suggesting a potential defect in the demarcation membrane system (Fig. 5.5B,D). Interestingly, emperipolesis was frequently seen in Nbeal2-/- MKs but not in WT MKs (Fig. 5.5E).

Figure 5.5. Abnormal ultrastructure of Nbeal2-/- bone marrow megakaryocytes. (A) Thin-section transmission electron micrographs of representative WT MKs show typical platelet territories (white arrowheads) and platelet-like structures containing α-granules. (B) In contrast, Nbeal2-/- MK are deficient in both α-granules and platelet territories (magnification X4000, scale bars = 2 μm). (C) Higher-magnification (X15 000) images of WT MK reveal multiple α-granules (white arrowheads) and platelet territories, whereas (D) α-granules (white arrowheads) are rare in Nbeal2-/- MK, which show poorly defined platelet territories (scale bars = 500 nm). (E) Example of the emperipolesis that was frequently observed in Nbeal2-/- MK; here, 3 exogenous cells (white arrowheads) are present in the MK (magnification X3000, scale bar = 10 μm).

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5.3.4 Absence of NBEAL2 leads to defects in megakaryocyte development and proplatelet formation

To understand the origins of thrombocytopenia in Nbeal2-/- mice, we first examined the development of Nbeal2-/- MKs. Native bone marrow cells were extracted and assessed for the population of CD41+ MKs over the course of a five-day culture. CD41+ cells were counted with confocal microscopy (Fig. 5.6A). Results revealed that although the proportion of CD41+ MKs were similar initially (Fig. 5.6B; WT: 0.228%േ0.094; Nbeal2-/-: 0.231%േ0.051), proportions of CD41+ MKs were significantly lower with Nbeal2-/- bone marrow cells after the five-day culture in comparison to WT (Fig. 5.6B; WT: 13.963%േ0.619; Nbeal2-/-: 7.675%േ0.119). Altogether, Nbeal2-/- MKs showed impairment in viability or development.

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Figure 5.6. Nbeal2-/- megakaryocytes showed reduced survival or differentiation. (A) Immunofluorescence imaging of a representative field of analysis. Bone marrow cells were stained with CD41 (Green) and DAPI. (B) Quantification of the percentage of CD41+ cells. Population of CD41+ cells increased by 70-folds over the course of 5-day culture while population of CD41+ cells derived from Nbeal2-/- mice were lower relative to that of the WT. 2500 cells were analyzed for the day 0 cultures per experiment per mice; n=3. Of the same mouse bone marrow cultures, 300 cells were analyzed for the day 5 cultures per experiment per mice; n=3. ** indicated P<0.01 with student’s t-test.

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Further studies were performed to determine the role of NBEAL2 in MK development. WT and Nbeal2-/- MKs were categorized into one of five developmental stages after a five-day culture and the populations of MKs at each stage were compared. CD41+ MKs were assigned to developmental stage denoted early (<20 μm in diameter), mature (>20 μm in diameter), extended (cells with membrane projections resembling early proplatelets), spread (cells with extensive membrane projections without proplatelets) or terminal proplatelet (extensive networks of cellular protrusions with emerging platelets). Representative confocal microscopy images of cells at each developmental stage categorized in this assay are shown in Fig. 5.7A. In this assessment, it was discovered that significantly higher proportions of Nbeal2-/- MKs were found in the early phase as well as in the spread phase, a developmental stage that likely precedes terminal proplatelets, than WT MKs (Fig. 5.7B). In conjunction with the evidence that Nbeal2-/- MKs are less likely to be found in proplatelet phase, these data suggest that Nbeal2-/- MKs are more likely to be stalled in the earlier phases of development and exhibited impairments in development. Further, DNA ploidies of native bone marrow MKs were assessed with flow cytometry. Results indicated a lower percentage of Nbeal2-/- MKs have DNA ploidies greater than 16N when compared to WT (Fig 5.8). Taken together, these data identified the requirement of NBEAL2 for MK development and maturation.

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Figure 5.7. Abnormal Nbeal2-/- megakaryocyte development. Immunofluorescence images of MKs cultured to the terminal proplatelet stage. (A) Cells present in fixed 5-day cultures were identified by immunofluorescence microscopy as MKs by size (>10 μm diameter), large/lobulated nuclei (light blue), and expression of lineage-specific CD41 (green) and VWF (red). Individual cells were classified by apparent developmental stage as early, <20 μm in diameter (ZY sections show these cells to be spheroidal); mature, >20 μm without projections (ZY sections show flattening with increased size); extending, round cells with membrane projections; spread, large cells with extensive membrane and tubulin (violet) projections but not showing clearly defined proplatelets; and terminal proplatelets emanate from these very large megakaryocytes. A distinctive pattern of nuclear retraction (light blue) and elaboration of nascent platelets defined by extensions of membrane (visualized by CD41, green) and cytoskeletal a-tubulin strands and loops (violet). (Extended focus; scale bar = 20 μm) (B) Population distributions of mean proportions of cells in each stage showed significant differences between Nbeal2-/- and WT (*P <.05; **P <.01, 2-tailed t test; n = 5 mice and 600 cells for both Nbeal2-/- and WT), with Nbeal2-/- cultures showing a markedly higher proportion of cells in both the early and spread stages and fewer in the mature, extending, and proplatelet stages.

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Figure 5.8. Delayed maturation of Nbeal2-/- megakaryocytes. Native bone marrow cells were stained with CD41 and propidium iodide then subjected to flow cytometry analysis. Population is normalized against the mode. Graph illustrates the distribution of Nbeal2-/- and WT CD41+ MK cells and ploidy, revealing reduced >16N Nbeal2-/- MKs (graph representative of 3 independent experiments).

The absence of α-granule cargo proteins in Nbeal2-/- platelets raised questions regarding the production and fate of these proteins (Fig. 5.2). Platelet precursor MKs were therefore investigated to understand the role of NBEAL2 in this process. Staining of permeabilized WT MKs showed a uniform distribution of VWF packages throughout the cell as expected (Fig. 5.9A). In contrast, while VWF is expressed in Nbeal2-/- MKs, VWF was primarily detected along the plasma membrane marked by CD41 (Fig. 5.9B). This VWF appeared to be extracellular as staining of non-permeabilized Nbeal2-/- MKs revealed strong signals of VWF lining the surface of the cell delineated by CD41 (Fig 5.9C). TEM analysis of Nbeal2-/- MKs with immunogold labelling confirmed the presence of VWF along the extracellular cellular surface, thus supporting previous findings (Fig. 5.9D). While VWF appears to be aberrantly

134 trafficked within mature MKs, this phenomenon was also evident during proplatelet formation. In WT MKs, VWF was packaged within the proplatelet buds of WT MKs as anticipated whereas VWF was clearly absent from the proplatelet buds of Nbeal2-/- MKs, and instead appeared to accumulate near the cell body (Fig. 5.10). Similar phenomena were seen in spread phase MKs (Fig. 5.11). Fig. 5.12 shows another representative example of a proplatelet phase MK with extensive extracellular VWF. These observations provide compelling evidences to indicate that VWF was lost in the MKs and was therefore unable to be trafficked towards the proplatelets. Determining the mechanism of VWF externalization will require further studies. We conclude that the lack of NBEAL2 leads to a premature loss of VWF as a result of aberrant trafficking.

Figure 5.9. Abnormal VWF distribution in Nbeal2-/- megakaryocytes. (A) Immunofluorescence imaging of a representative permeabilized mature extending stage WT megakaryocyte shows elaboration of demarcation membranes stained with CD41 (green), dispersed VWF expression (red), and a peripheral tubulin cytoskeletal meshwork (violet). (B) A representative Nbeal2-/- MK at the same stage shows a strong peripheral distribution of VWF, which is absent in Nbeal2-/- platelets; this MK also contains the nucleus of an exogenous cell in its cytoplasm visible in the XZ view of the merged confocal panel. Comparisons of non-permeabilized cells (C) confirm that VWF is abnormally concentrated near or on the surface of mature Nbeal2-/- MKs (scale bars in ZY panels = 5 μm; scale bars in XY, XZ, and extended focus panels = 10 μm). (D) Transmission electron micrograph of immunogold labeled VWF present on the surface of a Nbeal2-/- MK. Magnification X50 000; scale bar represents 500 nm.

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Figure 5.10. Von Willebrand factor is not trafficked to proplatelets in cultured Nbeal2-/- megakaryocytes. Immunofluorescence imaging of representative day 5 cultured WT and Nbeal2-/- MKs with proplatelets. Cells were stained with CD41 (green), von Willebrand factor (VWF; red) and α- tubulin (violet). VWF are seen in the proplatelet buds of WT MKs while it appears to be absent in the proplatelet buds of Nbeal2-/- MKs. Clusters of extracellular VWF are seen accumulating outside of the cell body of Nbeal2-/- MKs while this phenomenon is not observed in the WT MKs. (Extended focus; scale bar = 20 μm).

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Figure 5.11. Cultured Nbeal2-/- megakaryocyte in spread phase. Day 5 MKs in spread phase were imaged via immunofluorescence. A multilobed nucleus (light blue) is visualized with DAPI, cell membranes were illustrated with CD41 (green) alongside with α-tubulin (violet) for the visualization of cytoskeletal network. Clustered extracellular VWF (red) was detected along the plasma membrane. (Extended focus; scale bar = 20 μm).

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Figure 5.12. Cultured Nbeal2-/- megakaryocyte with proplatelets. MKs were cultured to the terminal proplatelet stage and imaged via immunofluorescence as described in Figure 5.8. A multilobed nucleus (light blue) is evident in the MK cell with membrane extensions visualized by CD41 (green) and the elaboration of nascent platelets defined by cytoskeletal α-tubulin strands and loops (violet). Extracellular von Willebrand factor (red) is also visible. (Extended focus; scale bar = 20 μm).

As indicated VWF was observed to be aberrantly released from the cell. What about other α- granule proteins? Upon examining TSP1, we noted that the level of intracellular TSP1 was reduced when compared to WT MKs (Fig. 5.13). Staining of proplatelets revealed that similar to VWF, TSP1 was absent in the proplatelet buds of Nbeal2-/- MKs and instead, appeared to accumulate near the cell body (Fig. 5.14). This suggested that similar to VWF, TSP1 was likely also released from the cell prior to proplatelet trafficking. Support of this hypothesis

138 comes from the analysis of Nbeal2-/- MK culture media revealing elevated levels of secreted TSP1 relative to WT MKs (Fig. 5.15). Levels of VWF were indistinguishable using this assay since VWF is an adhesive protein that was likely bound to the surface of MKs via GPIb as also on platelets during primary hemostasis (Peyvandi, Garagiola et al. 2011). Thus upon release from MKs, VWF readily adhered to the surface of MKs as seen in Fig. 5.9-5.12 and was therefore undetectable by the secretion assay. In contrast, TSP1 is less bound to MK cells and therefore more readily detected in the media. Taken together, these data indicate that the α-granule cargo proteins TSP1 and VWF were aberrantly released from Nbeal2-/- MKs.

Figure 5.13. Reduced TSP1 in Nbeal2-/- megakaryocytes. Immunofluorescence imaging of representative day 5 cultured WT and Nbeal2-/- MKs at mature and extend phases. Cells were stained with CD41 (green), TSP1 (red) and α-tubulin (violet). Nucleus visualized with DAPI. TSP1 signals are weaker in Nbeal2-/- MKs, suggesting a lower level of intracellular TSP1. (Extended focus; scale bar = 20 μm).

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Figure 5.14. TSP1 is not trafficked to proplatelets in cultured Nbeal2-/- megakaryocytes. Immunofluorescence imaging of representative day 5 cultured WT and Nbeal2-/- MKs with proplatelets. Cells were stained with CD41 (green), thrombospondin-1 (TSP1; red) and α-tubulin (violet). TSP1 is not observed in the proplatelets of Nbeal2-/- MKs and patches of extracellular TSP1 can be seen near the plasma membrane of the main cell body. (Extended focus; scale bar = 20 μm).

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Figure 5.15. TSP1 secretion by Nbeal2-/- megakaryocytes. Cultured MKs were purified on day 3 and incubated at equivalent concentrations in fresh media, from which samples were collected after 3 days and analyzed via immunoblot for thrombospondin 1 (TSP1) and transferrin as a sample loading control. The results indicate that unlike WT cells, Nbeal2-/- MKs secreted full-length TSP1 (FL-TSP1) at a level that was detectable against the background of full-length and cleaved forms present in the serum used to supplement the medium.

5.4 Discussion 5.4.1 NBEAL2 knockout leads to phenotypes that resemble human GPS in mice

In this study, we have shown that similar to human GPS, Nbeal2-/- mice displayed the hallmark features of macrothrombocytopenia, splenomegaly and platelet α-granule deficiencies (Nurden and Nurden 2007, Gunay-Aygun, Zivony-Elboum et al. 2010). In addition, Nbeal2-/- mice also displayed prolonged bleeding and their platelets had functional impairments. As these pathological phenotypes are the clinical manifestations of GPS, these findings support the suitability of Nbeal2-/- mice as a model for human GPS. Although human GPS is commonly associated with progressive bone marrow myelofibrosis, this phenotype was not detected in the bone marrows of Nbeal2-/- mice. GPS-associated bone marrow myelofibrosis is a progressive disease where the severity is positively correlated with age (Gunay-Aygun, Zivony-Elboum et al. 2010). It is therefore possible that four-months old mice used in our study have not yet developed myelofibrosis. It can also be possible that mice may naturally be more resilient to the development of myelofibrosis in the absence of NBEAL2.

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5.4.2 Nbeal2-/- platelets are functionally defective and contribute to bleeding in mice

In our study, we have shown that Nbeal2-/- platelets exhibited activation and aggregation defects (Fig. 5.4) These defects may lead to impairments in blood clot formation which may form the basis of the bleeding diathesis in Nbeal2-/- mice as observed. Despite the lack of α- granule cargoes including TSP1 and VWF in Nbeal2-/- platelets (Fig. 5.2), α-granule membrane constituent P-selectin was capable of surface mobilization during platelet activation (Fig. 5.3 and Fig. 5.4). This suggests that NBEAL2 is not essential for the secretion of α-granules and indicates that Nbeal2-/- platelets may have abnormal function due to the loss of α-granule cargo proteins. This is consistent with previous studies where ghost α- granules were described for GPS (Cramer, Vainchenker et al. 1985, Rosa, George et al. 1987, Maynard, Heijnen et al. 2010). Since in our Nbeal2-/- mice δ-granules in platelets were decreased, it is possible that the differential δ-granule release may contribute to the activation impairments of Nbeal2-/- platelets. However, as δ-granules are normal in GPS platelets, this rules out their contribution to the dysfunction of GPS platelets (Gerrard, Phillips et al. 1980). Taken together, these observations suggest that the bleeding diathesis in Nbeal2-/- mice is likely caused by the combination of impairment of platelet activation and lack of α-granule cargo proteins.

It is possible that other cell types may be affected by the loss of NBEAL2 which could contribute to the defects seen in Nbeal2-/- mice. While this possibility cannot be excluded, the predominant expression of NBEAL2 in hematopoietic cells (Albers, Cvejic et al. 2011), makes it likely that platelets are the primary contributors to the bleeding phenotypes seen in Nbeal2-/- mice. How NBEAL2 is involved in platelet activation is an outstanding question. It is possible that the different interactors of NBEAL2 may contribute to these functions. For example, 14-3- 3 has been shown to be central in mediating the signal transduction events of GPIb-IX-V- induced platelet activation (Chen, Ruggeri et al. 2018). Thus, it is possible that NBEAL2 may regulate platelet activity through its interaction with 14-3-3. NBEAL2 may also interact with FLNA to modulate platelet cytoskeleton to facilitate platelet shape change during activation (Nakamura, Stossel et al. 2011). Alternatively, NBEAL2’s interaction with SEC22B from the ER may provide platelets with the phospholipid membranes needed for shape change and spreading (Petkovic, Jemaiel et al. 2014).

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5.4.3 Nbeal2-/- platelets lack α-granule cargo proteins but retain membrane components of α-granules

ARC syndrome and GPS caused by mutations in VPS33B or VIPAS39 (VPS16B) and NBEAL2 respectively, are all characterized by bleeding and platelet α-granule deficiencies. In all, platelet α-granule cargo proteins are absent. Howerver, ARC platelets lack P-selectin while P-selectin remains present in Nbeal2-/- platelets, albeit at a lower level (Lo, Li et al. 2005, Urban, Li et al. 2012). We have shown in our study that NBEAL2 is not essential for P- selectin mobilization during platelet activation (Fig. 5.3 and Fig. 5.4). This is consistent with previous findings where it was shown that P-selectin membranes in GPS platelets were secretion competent (Rosa, George et al. 1987). These data suggest the likely presence of immature α-granules in Nbeal2-/- platelets that could be partially functional. Immunogold analysis of GPS platelets has shown that P-selectin is localized to the limiting membranes of vacuolar structures (Rosa, George et al. 1987). Most of these P-selectin vacuoles are empty but some were labelled positive for the presence of α-granule cargo protein albumin, suggesting the nature of these empty vacuoles are likely α-granules that failed to mature. This is consistent with our ultrastructure analysis of Nbeal2-/- MKs and platelets where an abundance of vacuolated structures were found (Fig. 5.1 and Fig. 5.5). These phantom granules are similar in size to α-granules and are scantly labelled with α-granule proteins VWF and albumin, suggesting that they represent immature α-granules (Cramer, Vainchenker et al. 1985, Rosa, George et al. 1987). From these observations, it is likely that NBEAL2 is not essential for the initial formation of α-granule precursor vesicles but may instead be required for the trafficking of α-granule cargoes into a pre-existing immature α- granule. In contrast, ARC platelets lack P-selectin. This suggest that VPS16B/33B may be responsible for generating the α-granule precursor vesicles where NBEAL2 then acts to promote the maturation of immature α-granules via α-granule cargo protein trafficking.

5.4.4 NBEAL2 may be required for the stabilization of α-granules

Based on the above considerations, it is unlikely that NBEAL2 is required for the initial formation of immature α-granules. Instead NBEAL2 is likely involved in the trafficking of α- granule cargo proteins. Immunofluorescence and immunogold studies of Nbeal2-/- MKs have revealed the aberrant externalization of α-granule protein VWF (Fig. 5.9). As α-granule deficiency was detected in maturing Nbeal2-/- MKs by ultrastructural imaging (Fig. 5.5), indicates that α-granule cargo protein loss occurred in MKs prior to platelet production. The α- granule cargo protein loss from MKs seen in Nbeal2-/- MKs may contribute to the

143 development of progressive bone marrow fibrosis where the persistent release of growth factors could stimulate the migration and proliferation of fibroblasts into the bone marrow of GPS patients. While it is evident that NBEAL2 is involved in the trafficking of α-granule proteins in MKs, the details of NBEAL2 function have not been established. Nonetheless, our studies show that NBEAL2 is required for retaining MK α-granule cargo proteins such as VWF and TSP1 in the intracellular compartments (Fig. 5.9-Fig. 5.15). The reduction of P- selectin level in Nbeal2-/- platelets suggest that NBEAL2 may also be responsible for maintaining the stability of α-granules, which may in turn facilitate the retention of α-granule proteins. Since we have previously shown that NBEAL2 localizes primarily to P-selectin in primary MKs (Fig. 3.10), it is reasonable to propose that NBEAL2 functions in the stabilization of α-granules and thereby prevents the aberrant secretion of α-granule proteins. Further analysis of α-granule protein trafficking in Nbeal2-/- MKs should help to elucidate the role of NBEAL2 for α-granule biogenesis.

5.4.5 NBEAL2 is required for normal megakaryocyte development and platelet production

GPS patients and Nbeal2-/- mice have thrombocytopenia. The cause of the thrombocytopenia is however unknown. Platelet destruction is not the primary cause of thrombocytopenia as WT and Nbeal2-/- platelets exhibit similar life span in-vivo (Deppermann, Cherpokova et al. 2013). To examine potential platelet production defects, Nbeal2-/- MKs were analyzed. Nbeal2-/- MK showed delayed maturation (Fig. 5.7), decreased survival (Fig. 5.6), decreased ploidy (Fig. 5.8), and developmental abnormalities such as underdeveloped platelet territories or DMS (Fig. 5.5). Ultimately, the combined effects of these cellular abnormalities likely contribute to the reduction of proplatelet production by Nbeal2-/- MKs (Fig. 5.7), thereby leading to thrombocytopenia.

As with other BDCPs, NBEAL2 likely functions as a scaffold protein and may be involved in multiple cellular processes through different protein-protein interactions. Some of these may be required for MK maturation that do not involve α-granule biogenesis. Using AP-MS we have identified a number of potential (Table 4.3) NBEAL2 binding proteins some of which could facilitate different cellular events. As 14-3-3 is implicated in cell survival, development and growth (Dong, Kang et al. 2007), NBEAL2-14-3-3 interactions may be important in mediating the growth, survival and development of MKs. FLNA is a protein that is involved in the remodeling of cortical actin cytoskeleton (Nakamura, Stossel et al. 2011). As an actin-

144 based motor force is an important driver of proplatelet branching (Patel, Hartwig et al. 2005) and FLNA-deficient mice also displaying macrothrombopenia and abnormal DMS development similar to Nbeal2-/- mice (Begonja, Pluthero et al. 2015), the interaction between NBEAL2 and FLNA may be important for DMS development and platelet production. SEC22B is an ER protein that has been shown to be important for the expansion of PM and phagosome maturation in dendritic cells (Cebrian, Visentin et al. 2011, Petkovic, Jemaiel et al. 2014). Proplatelet production by MKs requires a large supply of membranes which is derived from the DMS. Multiple membrane trafficking pathways likely occurr in MKs during proplatelet production in order to meet the intense demands of generating platelets. MCS between the ER and organelles has been shown to be a key component in non-vesicular lipid and membrane transfer (Toulmay and Prinz 2011, Lev 2012), as is the case of SEC22B during PM expansion (Petkovic, Jemaiel et al. 2014). It is possible that NBEAL2 and SEC22B forms MCS, thereby allowing the transfer of membranes from the ER to the PM to support proplatelet production. SEC22B has also been implicated in the maturation of phagosomes and the formation of MCS during endosome maturation (Cebrian, Visentin et al. 2011, Friedman, Dibenedetto et al. 2013). MCS between ER resident SEC22B and NBEAL2, which we have shown colocalizes with P-selectin (Fig. 3.10), may promote the maturation of α- granules. While plausible, the functional significance of these NBEAL2 interactors will require further investigation. Nevertheless, our study here has identified that NBEAL2 is required for megakaryocyte survival, development, platelet production and α-granule protein trafficking.

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Chapter 6

Role of NBEAL2 in α-granule cargo trafficking

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6.1 Abstract

Platelets are made by MKs, and to elucidate the role of NBEAL2 in α-granule formation we investigated intracellular trafficking of both megakaryocyte-synthesized (VWF) and endocytosed (FGN) α-granule cargo in WT and Nbeal2-/- mouse MKs. Their initial uptake of labeled FGN was similar, but it was released by Nbeal2-/- megakaryocytes and retained by WT cells. Tracking experiments revealed sequential colocalization of endocytosed fibrinogen with compartments expressing RAB5, RAB7 and P-selectin in WT and Nbeal2-/- MKs, however, FGN and VWF appeared to be externalized by RAB11 recycling endosomes in Nbeal2-/- cells. Previously we have shown the colocalization and coimmunoprecipitation of NBEAL2 with P-selectin in human megakaryocytes and platelets, indicating that NBEAL2 associates with α-granules. Taken together, we conclude that NBEAL2 is required for the retention of endocytosed and megakaryocyte-synthesized α-granule cargo proteins.

6.2 Introduction

We have observed that NBEAL2 is required for normal megakaryocyte development and α- granule production, and that in mouse MKs a lack of NBEAL2 results in the aberrant release of the granule cargo proteins VWF and TSP1. The latter observation indicates that NBEAL2 may be involved in the trafficking of α-granule proteins and/or their retention by maturing granules. Intracellular trafficking of α-granules and their cargo has not been extensively studied, primarily owing to the difficulties associated with the experimental manipulation of MKs. FGN is endocytosed by MKs (Handagama, Rappolee et al. 1990, Louache, Debili et al. 1991), providing the opportunity to track protein into cells and along transport pathways associated with trafficking of α-granule cargo proteins. In this study we examined FGN trafficking in WT and Nbeal2-/- mouse MKs to reveal the role of NBEAL2 in α-granule biogenesis.

6.3 Results 6.3.1 Aberrant α-granule cargo protein trafficking in Nbeal2-/- megakaryocytes

In the previous chapter we observed that the MK-synthesized α-granule cargo proteins VWF and TSP1 were externalized from MKs in the absence of NBEAL2. Murine and human MKs are incapable of synthesizing FGN, hence they are dependent on endocytosis from plasma to obtain FGN protein for packaging into α-granules (Handagama, Rappolee et al. 1990, Louache, Debili et al. 1991). We have previously reported that compared to WT, platelets

147 from Nbeal2-/- mice have significantly reduced levels of FGN, VWF, TSP1 and P-selectin (Kahr, Lo et al. 2013). Immunoblot analysis confirmed these observations for platelets and MKs from the animals used in this study, together with the absence of NBEAL2 expression by Nbeal2-/- mice (Fig. 5.2).

In previous studies we observed that VWF synthesized by Nbeal2-/- MKs is abnormally trafficked to the cell surface and externalized (Fig. 6.1), and since the FGN level in Nbeal2-/- platelets is also reduced, this suggests that NBEAL2 may be involved in the trafficking of MK- synthesized and endocytosed granule cargo (Fig. 6.2). Using immuno-electron microscopy we confirmed that while VWF and FGN associate with α-granules in native bone marrow MKs from WT mice, these proteins show no association with subcellular structures in Nbeal2-/- MKs (Fig. 6.3), consistent with aberrant α-granule cargo protein trafficking in Nbeal2-/- MKs.

Figure 6.1. Abnormal VWF trafficking in Nbeal2-/- megakaryocytes. Cultured MKs stained for CD41 (green), tubulin (magenta) and VWF (red; blue = DAPI staining of DNA) and imaged by confocal fluorescence microscopy show internalized VWF in WT cells at early, mature and proplatelet-forming (Extended) stages, and extensive VWF externalization in late-stage Nbeal2-/- MKs (extended focus images; bars = 20 µm).

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Figure 6.2. FGN is reduced in Nbeal2-/- platelets. (Left panel) Nbeal2-/- platelets were lysed and analyzed for the level of FGN. Immunoblot results indicate a lower level of FGN in Nbeal2-/- platelets, suggesting a requirement of NBEAL2 in FGN trafficking. (Right panel) Cultured bone marrow MKs were grown in serum-free media and analyzed for the presence of FGN. As the serum-free media does contain FGN, the cultured MKs also do not harbor FGN.

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Figure 6.3. Abnormal VWF and FGN localization in Nbeal2-/- megakaryocytes. Native bone marrow MKs stained with immunogold-labeled FGN or VWF, and examined via transmission electron microscopy show that in WT cells most of the FGN and VWF localized in or near electron-dense α- granules (inset arrowheads), while in Nbeal2-/- MKs both proteins appeared to be randomly distributed throughout the cell. Bars = 500 nm for FGN (60,000X magnification) and 100 nm for VWF (80,000X magnification); insets are magnified 2X further.

6.3.2 Expression of fluorescent-tagged NBEAL2 in Nbeal2-/- mice.

In GPS patients and Nbeal2-/- mice the loss of NBEAL2 expression is ubiquitous, but it is generally assumed that the GPS phenotype is attributable to the loss of NBEAL2 expression in MKs. To examine this hypothesis, we constructed a Flag-GFP-NBEAL2 controlled by the promoter for the megakaryocyte-specific gene ITGA2B, which encodes integrin alpha 2b (aIIb). Transgenic introduction of this construct into Nbeal2-/- mice will: 1) allow us to determine whether rescue of NBEAL2 expression in MKs is sufficient to reverse the GPS phenotype in Nbeal2-/- mice, and 2) track fluorescently-tagged NBEAL2 within MKs and platelets via live imaging, providing insights into its cellular role.

Linearized aIIb-Flag-GFP-NBEAL2 DNA was microinjected into Nbeal2-/- mouse embryos. Transgenic mice were then bred and assayed for the integration of the transgene through PCR DNA analysis. Platelets from the positive mice were further examined for the expression of Flag-GFP-NBEAL2 via immunoblot, which confirmed expression of transgenic NBEAL2 in

150 platelets from the first generation of transgenic mice (Fig. 6.4). Mice that tested positive for the expression of the transgene were bred for three generations. When platelets from the fourth generation were again assayed for expression of transgenic NBEAL2 it was unexpectedly not detected when blots were probed with either anti-Flag (Fig. 6.5A) or anti- NBEAL2 antibodies (Fig. 6.5B). Earlier generations of transgenic mice were tested, and the results indicated that the transgene was not expressed after the first generation (Fig. 6.6). The cause of this loss of expression was not known.

Figure 6.4. Transgenic expression of GFP-NBEAL2 in Nbeal2-/- mice. αIIb-Flag-GFP-NBEAL2 was introduced into Nbeal2-/- mice via embryo microinjections. Platelets from the first generation of transgenic mice were analyzed by immunoblot for the expression of the transgene. Of 7 mice examined, platelets from 6 showed expression of Flag-GFP-NBEAL2. These mice were crossbred to maintain and expand the population.

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Figure 6.5. The expression of the NBEAL2 transgene in Nbeal2-/- mice did not propagate. Nbeal2- /- mice expressing the Flag-GFP-NBEAL2 transgene were bred for several generation and platelets from the fourth generation were analyzed again for the expression of the transgene. (A) Immunoblot was probed with Flag antibody. A weak protein signal was observed with the transgenic mice platelets that may resemble Flag-GFP-NBEAL2 based on the predicted molecular weight. (B) Immunoblot was then probed with NBEAL2 antibody. In this case, the expression of the transgene was not observed. The results suggest that the transgene expression did not propagate through the generations of mice.

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Figure 6.6. Validating the expression of NBEAL2 in Nbeal2-/- mice. Platelets from the second generation of mice were analyzed for the expression of Flag-GFP-NBEAL2. Immunoblots were probed for NBEAL2 and GFP. The results indicate the lack of expression of the transgene.

6.3.3 Intracellular fibrinogen is significantly reduced in Nbeal2-/- megakaryocytes following endocytosis

We initially examined FGN uptake by WT and Nbeal2-/- mouse MKs cultured in serum-free and thus FGN-free medium by incubating cells with FGN conjugated with Alexa Fluor 488 for 24 h, followed by fixation and imaging via confocal fluorescence microscopy. As expected, these cultured cells do not express endogenous FGN (Fig. 6.2). The results (Fig. 6.7) show that both WT and Nbeal2-/- MKs took up FGN, but while WT cells typically showed abundant endocytosed FGN localized to defined cellular structures resembling MVBs/late endosomes, Nbeal2-/- MKs contained markedly less endocytosed FGN distributed among indistinct compartments. The identification of the large structures associated with endocytosed FGN in WT MKs as MVBs/late endosomes was confirmed via co-staining with RAB7 (Fig. 6.8).

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Figure 6.7. Fibrinogen endocytosis by WT and Nbeal2-/- megakaryocytes. Purified day 4 MKs were incubated with 0.1 mg/mL labelled fibrinogen (green) for 24 h prior to fixation and staining for P-selectin (magenta), CD41 (red) and nuclear DNA (blue). Confocal fluorescence microscopy images of representative cells show reduced endocytosed FGN in Nbeal2-/- MKs compared to WT, and also a different distribution pattern with FGN in WT cells associated with structures resembling multivesicular bodies (extended focus images, bars = 10 µm).

Figure 6.8. Endocytosed fibrinogen is associated with multivesicular body-like structures in WT megakaryocytes. Confocal fluorescence microscopy images of WT bone marrow MKs incubated with Alexa Fluor 488 labeled FGN for 24 h shows association of endocytosed FGN (green) with RAB7- positive (magenta) structures resembling late endosomes/multivesicular bodies (inset). (Confocal Z- sections; scale bar = 10 µm).

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6.3.4 Endocytosis is not affected by the loss of NBEAL2

To assess FGN uptake, WT and Nbeal2-/- MKs were incubated with fluorescent FGN for 30 min or overnight (O/N), then imaged with confocal microscopy. Representative images are shown in Fig. 6.9A. Results indicate that fluorescent FGN signals are similar in WT and Nbeal2-/- MKs that undergo a short incubation (30 min) with fluorescent FGN. However, the fluorescent FGN signals are lower in Nbeal2-/- MKs with O/N incubation. The differential FGN uptake by WT and Nbeal2-/- MKs was quantified by incubating bone marrow derived MKs with fluorescent FGN for 1 or 24 h, and then assessing FGN content via flow cytometry. The results (Fig. 6.9B-C) indicate that after 1 h incubation WT and Nbeal2-/- MKs showed little difference in fluorescent signal from endocytosed FGN, but after 24h the signal was significantly weaker for Nbeal2-/- cells. These results indicate that while the initial rate of FGN uptake is similar for WT and Nbeal2-/- MKs, the latter show markedly reduced accumulation of endocytosed protein.

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Figure 6.9. Different fibrinogen endocytosis by WT and Nbeal2-/- megakaryocytes. Bone marrow derived cultures were incubated with FGN conjugated with Alexa Fluor 488 prior to analysis via flow cytometry or confocal fluorescence microscopy. (A) Representative images show similar labeled FGN (green) uptake in WT and Nbeal2-/- MKs after 30 min incubation (upper), and higher FGN content in WT cells after O/N incubation (Z-sections; scale bars = 10 µm). (B) Bone marrow derived cultures were incubated with labelled FGN (FGN-Alexa Fluor 488) for 1 or 24 h (O/N) prior to flow cytometry analysis of MKs for green fluorescence intensity. At both time points, WT and Nbeal2-/- MKs show increased endocytosed protein signal relative to cells incubated without labelled FGN (Control). After 1h incubation there is little difference between WT and Nbeal2-/- MKs in signal expressed as median fluorescence intensity (MFI), but after 24h, Nbeal2-/- cells have a significantly weaker signal. Data representative of 3 independent experiments; error bars = standard deviation, * = P<0.05 via one-way ANOVA with Tukey post-test, NS = P>0.05.

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6.3.5 Fibrinogen is not lost as a result of lysosomal degradation in Nbeal2-/- megakaryocytes

The possibility that endocytosed FGN is differentially degraded by WT and Nbeal2-/- MKs was examined by incubating purified MKs with biotinylated FGN (bio-FGN) for 24 h and measuring it in cell lysates via protein blotting and probing with HRP conjugated streptavidin (Fig. 6.10A- B). The results confirmed reduced accumulation of endocytosed FGN by Nbeal2-/- MKs, and also showed no indication of FGN degradation in cell lysates. A protease-treated sample was used as a positive protein degradation control (lane 2, Fig. 6.10A). Treatment with chloroquine, which inhibits lysosomal protein degradation (Seglen, Grinde et al. 1979), had no effect (lanes 5 & 6 of Fig. 6.10A and Fig. 6.10B). Thus the reduced accumulation of endocytosed FGN observed in Nbeal2-/- MKs cannot be attributed to proteolytic degradation, consistent with the reported absence of secondary lysosomes in MKs (Bentfeld-Barker and Bainton 1982).

Figure 6.10. Endocytosed fibrinogen is not aberrantly degraded in Nbeal2-/- megakaryocytes. (A) Probing with streptavidin-HRP of lysates isolated from purified MKs after 24 h incubation with biotinylated FGN indicates significantly less endocytosed protein present in Nbeal2-/- cells compared to WT. (B) Densitometric analysis of ratios of mean density of FGN bands relative to GAPDH confirms a significant difference (* = P<0.05 one-way ANOVA with Tukey post-test; NS = P>0.05; bars = standard deviation; data representative of 3 independent experiments). The same results show no indication of protein degradation in MKs; a sample where biotinylated FGN was incubated with proteinase K is shown for comparison (FGN+protease; *denotes the same degraded FGN band in upper and lower panels). Chloroquine treatment of MKs (WT+CQ, Nbeal2-/- + CQ) did not affect FGN uptake or degradation.

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6.3.6 Absence of NBEAL2 results in aberrant secretion of fibrinogen

The retention of FGN by WT and Nbeal2-/- MKs was examined by incubating cells with bio- FGN for 24 h, transferring them to fresh media lacking FGN, and collecting media samples at 0, 4, 8 and 24 h time points for concentration and protein content analysis. The results (Fig. 6.11A) indicate a more rapid release of FGN from Nbeal2-/- MKs relative to WT cells. The medium supplement transferrin was used as a sample loading control (Fig. 6.11A, bottom panel). The difference was not due to differential cell death/lysis, since probing of cell releasates for GAPDH was negative (not shown). Probing of cell lysates derived from MKs at the initial and final time points (0 and 24 h, Fig. 6.11B) confirmed greater loss of bio-FGN from Nbeal2-/- MKs. Densitometric analysis confirmed that the release of bio-FGN from WT and Nbeal2-/- MKs was significantly different (Fig. 6.11C). A similar differential pattern of release over a 3-day period was observed for MK-synthesized TSP1 (Fig. 5.15). Taken together, these results suggest that NBEAL2 is not involved in FGN endocytosis by MKs, but it appears to be required for the retention of both endocytosed and MK-synthesized α-granule cargo.

Figure 6.11. Greater release of endocytosed fibrinogen by Nbeal2-/- megakaryocytes. (A) MKs incubated with biotinylated FGN for 24 h were washed and transferred to fresh medium, from which samples were collected at 4 h intervals and analyzed for released biotinylated FGN, plus transferrin present in the medium. (B) Lysates from cells sampled 0 and 24 h time points show decreased retention of endocytosed FGN in Nbeal2-/- MKs relative to WT. (C) Densitometric analysis shows an increased rate of FGN release by Nbeal2-/- MKs relative to WT (* = P<0.05 one-way ANOVA with Tukey post-test; NS = P>0.05; bars = standard deviation; data representative of 3 independent experiments).

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6.3.7 Fibrinogen is not retained in P-selectin positive vesicles and is secreted via Rab11+ vesicles in the absence of NBEAL2

The role of NBEAL2 in the intracellular trafficking of endocytosed FGN by MKs was examined by monitoring the passage of labeled protein through vesicular compartments using fluorescence microscopy. Previous studies reported that MKs use a classic endocytic pathway to traffic FGN to α-granules, thus we focused on endosomes associated with this pathway that were identified via the markers RAB5 (early endosomes), RAB7 (late endosomes/MVBs), P-selectin (nascent/mature α-granules) and VWF (mature granules). The possibility that FGN trafficking is grossly abnormal in Nbeal2-/- MKs made it necessary to examine the potential involvement of other compartments, including endosomes associated with rapid recycling (RAB4 positive), slow recycling and exocytosis (RAB11 positive) (Ward, Martinez et al. 2005, Sugawara, Shibasaki et al. 2009, Takahashi, Kubo et al. 2012). We also examined the possible involvement of trafficking and secretion via the recycling transferrin receptor (TFR), exosomes from MVBs involving RAB27A and RAB27B (Ostrowski, Carmo et al. 2010), the VAMP2-associated compartment involved in regulated exocytosis in granulocytes (Lacy, Logan et al. 2001, Mollinedo, Martin-Martin et al. 2003) and lysosomes (LAMP1).

An initial analysis of colocalization of endocytosed FGN with these markers was done for day 4 WT MKs incubated for 24 h with labeled FGN. The images shown in Fig. 6.12A are representative of those used for a quantitative colocalization analysis based on Pearson’s correlation coefficient (Fig. 6.12B). This analysis indicated substantial differences in the intracellular destination of endocytosed FGN in WT and Nbeal2-/- MKs. The expected relatively strong association of FGN with P-selectin and VWF seen in WT MKs was not observed in Nbeal2-/- MKs, where the strongest FGN association was with the RAB11+ endosomal compartment.

To more precisely assess the intracellular trafficking of FGN in MKs, we incubated cells with fluorescently labeled FGN for 30 min, after which they were transferred to semi-solid Matrigel culture on coverslips. Samples were fixed after a further 1, 2, 4, 6 or 8 h, and then samples from all time points were stained for one of RAB4, RAB5, RAB7, RAB11, P-selectin or LAMP1, and the association of endocytosed FGN with these markers was assessed via confocal fluorescence microscopy. Representative stained images for RAB5, RAB7, P- selectin and RAB11 are shown in Fig. 6.13A-D. To quantify the colocalization of FGN with

159 these vesicle markers we used the Manders overlap coefficient (Fig. 6.13E). The results of this analysis indicate that FGN first associates (0 h) with RAB5-positive early endosomes in WT and Nbeal2-/- MKs. Within 1 h, WT and Nbeal2-/- MKs also showed an increased association of FGN with RAB7+ endosomes (Fig. 6.13E). In WT MKs this was followed by an increasing association of FGN with P-selectin. In contrast, FGN in Nbeal2-/- cells showed a peak association with P-selectin at 2 h that rapidly declined as association with RAB11+ endosomes increased, peaking at 6 h. After 8 h, most of the FGN endocytosed by WT MKs was associated with P-selectin, while the low amount retained by Nbeal2-/- MKs showed little detectable association with any vesicular marker. Thus in contrast to WT cells, Nbeal2-/- MKs traffic FGN partially and/or briefly through the endocytic pathway leading to nascent α- granules and then most of the endocytosed protein passes to RAB11+ recycling endosomes and is secreted.

This conclusion was supported by the results of experiments where purified day 3 MKs were sequentially incubated for 24 h each with green or red fluorescently-tagged FGN. Images of these cells (Fig. 6.14) showed considerable convergence of both forms of labeled FGN in WT MKs and their association with P-selectin. In contrast, these associations were much less evident in Nbeal2-/- MKs. Taken together these results indicate that NBEAL2 is required for P- selectin positive vesicles to acquire and/or retain FGN and other granule cargo proteins.

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Figure 6.12. Endocytosed fibrinogen is abnormally distributed in Nbeal2-/- megakaryocytes. (A) Purified MKs were incubated with 0.1 mg/mL of Alexa Fluor 488 labeled FGN for 24h on day 4, then fixed and stained for RAB5, RAB4, RAB7, RAB11, RAB27A, RAB27B, P-selectin, TFR, VWF, LAMP1 or VAMP2. Images shown are representative (Z-sections; scale bars = 10 µm) of those used to calculate Pearson’s correlation coefficient values indicating colocalization of FGN with these markers. (B) A graph of the resulting mean values (bars = standard deviation) indicates a relatively strong association of FGN with P-selectin and VWF in WT MKs that is not seen in Nbeal2-/- cells, which show a unique strong association of FGN with RAB11 (n=3 independent experiments with 15 cells for each condition; *P<0.05; **P<0.01, Mann-Whitney U test).

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Figure 6.13. Different trafficking of endocytosed fibrinogen by WT and Nbeal2-/- megakaryocytes. Day 5 cultured WT and Nbeal2-/- MKs were incubated with fluorescent FGN (green) for 30 minutes, then washed and either fixed and centrifuged onto coverslips (0 h samples) or seeded onto Matrigel- coated coverslips and fixed after 1, 2, 4, 6 or 8 h. Fixed cells were stained and imaged to assess the association of endocytosed FGN with markers of endosomes (RABs 4, 5, 7,11; P-selectin) and lysosomes (LAMP1). Representative Z-section images are shown for cells stained for A-D: RAB5, RAB7, P-selectin, RAB11 (bars = 10um; boxed areas magnified in insets). Colocalization of endocytosed FGN with endosomal/lysosome markers was quantified in images of cells >20 µm diameter via the Manders overlap coefficient. E, Graphs show mean values (bars = standard deviation) from 3 experiments with n = 15-25 cells at each time point for WT and Nbeal2-/- cells; *=P<0.05 for inter-strain comparisons (Kruskal-Wallis test with Dunn’s post-hoc test). Results indicate an early association of endocytosed FGN with RAB5 associated early endosomes in WT and Nbeal2-/- MKs, with both also showing a shift of FGN towards RAB7 associated endosomes after 1-2 h. In WT MKs this is followed by an increasing association of FGN with P-selectin from 2 h on, while FGN in Nbeal2-/- cells shows a peak association with P-selectin at 2 h, followed by a steady decline as FGN appears in RAB11 associated endosomes after 6 h. After 8 h most of the FGN endocytosed by WT MKs is associated with P-selectin, while the much lower level retained by Nbeal2-/- MKs shows no definite endosomal association.

Figure 6.14. Convergence of endocytosed fibrinogen in WT megakaryocytes. Purified day 3 MKs were incubated with 0.1mg/mL of FGN tagged with Alexa Fluor 488 (Fibrinogen-488; green) for 24h, washed, incubated with the same concentration of FGN tagged with Alexa Fluor 555 (Fibrinogen-555; red) for 24h, then washed, fixed and immunostained for P-selectin (magenta). These representative confocal fluorescence microscopy images (mid-cell Z-sections, bar = 10 µm) show that in WT MKs both forms of endocytosed FGN converged on the same compartments, which were associated with P- selectin. FGN convergence and association with P-selection was much less evident in Nbeal2-/- MKs.

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6.3.8 VWF is aberrantly trafficked to Rab11+ vesicles in Nbeal2-/- megakaryocytes

The results obtained for endocytosed FGN led us to examine the possibility that the abnormal trafficking of VWF observed in Nbeal2-/- MKs (Kahr, Lo et al. 2013) may also involve the RAB11-associated recycling endosomal compartment. Examination of mature MKs stained for RAB11 and VWF (Fig. 6.15A-B) showed extensive colocalization in Nbeal2-/- cells, but not in WT MKs. A similar association was observed in purified MKs incubated with fluorescent FGN for 24 h, then placed onto poly l-lysine-coated coverslips, fixed and stained for RAB11 and VWF. Most Nbeal2-/- MKs showed colocalization of RAB11 with FGN or VWF, and in some cells all 3 proteins were observed to colocalize (Fig. 6.16). Thus in the absence of NBEAL2, RAB11+ recycling endosomes appear to be involved in the abnormal trafficking and secretion of both endogenously-synthesized and endocytosed α-granule cargo proteins in MKs.

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Figure 6.15. Abnormal trafficking of VWF by Nbeal2-/- megakaryocytes. (A) Representative confocal fluorescence microscopy images of mature day 5 MKs cultured in matrigel fixed and stained for RAB11 and MK-synthesized VWF show strong colocalization in Nbeal2-/- but not WT MKs. (B) Manders overlap coefficient analysis showing significant higher association between VWF and RAB11 in Nbeal2-/- MKs compared to WT MKs (n=3 independent experiments with 15-25 cells for each condition; *P<0.05, Mann-Whitney U test).

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Figure 6.16. Abnormal trafficking of endogenously-synthesized and endocytosed granule cargo by Nbeal2-/- megakaryocytes. Day 4 MKs were purified, incubated with fluorescent FGN for 24 hours, then placed on polylysine-coated coverslips, fixed, stained and imaged. Strong colocalization of VWF, FGN and RAB11 was observed in Nbeal2-/- MKs, but not in WT cells (Z-sections; scale bars = 10 µm).

6.3.9 Nbeal2-/- megakaryocytes do not exhibit premature α-granule release

P-selectin is mobilized to the surface of MKs and platelets during α-granule release (Fig. 5.3C-D), thus if the loss of granule cargo from Nbeal2-/- MKs is associated with premature α- granule release, they should show surface-mobilized P-selectin. Staining of non- permeabilized cells showed that the expected detection of released VWF on Nbeal2-/- MKs was not accompanied by surface mobilization of P-selectin (Fig. 6.17A), which was detected on the surface of thrombin-stimulated WT cells. These results indicate that VWF release by Nbeal2-/- MKs is not due to premature release from α-granule or other vesicles containing P- selectin. RAB11 was detected along with VWF on the surface of Nbeal2-/- MKs (Fig. 6.16B), confirming that loss of VWF and other granule cargo is likely mediated by RAB11+ vesicles.

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Figure 6.17. Release of VWF by Nbeal2-/- megakaryocytes is not mediated by P-selectin+ vesicles. (A) Nbeal2-/- and WT MKs were fixed and immunostained without permeabilization for CD41 (green), VWF (red) and P-selectin (violet); also tested were WT MKs treated with thrombin (1 unit/mL for 15 minutes) as an activation control. P-selectin was only detected on the surface of activated WT MKs, which showed surface VWF along with Nbeal2-/- MKs. (B) MKs were stained for CD41 (violet), VWF (red) and RAB11 (green), which were all observed near the plasma membrane in Nbeal2-/- MKs, while VWF and RAB11 were not seen on WT MKs. These results indicate that premature α-granule release is unlikely to be a major cause of VWF release in Nbeal2-/- MKs, which appears to be mediated by RAB11+ vesicles.

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6.4 Discussion

6.4.1 Absence of NBEAL2 results in the aberrant loss of α-granule cargo proteins in megakaryocytes

We have shown that the MK-synthesized α-granule cargo proteins VWF and TSP1 are lost from Nbeal2-/- MKs (Figure 6.1), and previous studies have observed VWF deposition near the DMS of cultured GPS MKs via electron microscopy (Breton-Gorius, Vainchenker et al. 1981, Drouin, Favier et al. 2001). VWF and TSP1 are almost absent from Nbeal2-/- platelets (Fig. 5.2A), which show somewhat higher levels of FGN (Fig. 6.2). Since platelets have been reported to be capable of endocytosing fibrinogen (Huang, Joshi et al. 2016), it was not possible to determine whether FGN is aberrantly trafficked in MKs without directly examining this aspect of α-granule development. Here, using a range of techniques including immunofluorescence and immunogold microscopy and secretion assays, we have demonstrated that like VWF and TSP1, FGN is aberrantly trafficked by and released from Nbeal2-/- MKs.

6.4.2 Attempted rescue of Nbeal2-/- mice with fluorescent NBEAL2

In our previous study we characterized the Nbeal2-/- mouse and revealed that NBEAL2 is required for the formation of α-granules and normal development of MKs (Kahr, Lo et al. 2013). A potential caveat to these studies is that in Nbeal2-/- mice NBEAL2 deficiency is not limited to MKs, and this protein is expressed in other hematopoietic cells that may influence the GPS phenotype. In an attempt to address this possibility we tried to rescue MK expression of NBEAL2 in Nbeal2-/- mice using constructs where human NBEAL2 is tagged by a fluorescent probe to potentially facilitate intracellular protein imaging. While initial results were promising, we were not able to generate rescued mice. Possible explanations include loss or inactivation of the NBEAL2 transgene, or the incompatibility of human NBEAL2 in mice. Nevertheless, since Nbeal2-/- MKs grown in culture exhibit the same α-granule protein loss evident in Nbeal2-/- mice (Fig. 6.11 and Fig. 6.13), we are confident that NBEAL2 plays a critical role in MK α-granule biogenesis. Recent reports indicate that NBEAL2 may also be important in mast cells and neutrophils (Drube, Grimlowski et al. 2017, Sowerby, Thomas et al. 2017).

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6.4.3 Loss of fibrinogen in Nbeal2-/- megakaryocytes is not caused by aberrant endocytosis or degradation

The low FGN levels observed in Nbeal2-/- MKs and platelets (Fig. 6.2 and Fig. 6.7) could arise due to one or more of: defective FGN endocytosis, increased FGN degradation, and aberrant trafficking/secretion by MKs. Since MKs derived directly from marrow would be expected to be fully-loaded with FGN in vivo, in order to examine FGN endocytosis and trafficking we cultured bone marrow-derived MKs in FGN-free media so that they lacked FGN (Fig. 6.2). Flow cytometry analysis of FGN uptake by cultured MKs showed that endocytosis is not affected by the absence of NBEAL2 (Fig. 6.9 B-C), consistent with evidence that surface levels of FGN receptor required for endocytosis (Handagama, Bainton et al. 1993, Handagama, Scarborough et al. 1993) are normal in Nbeal2-/- MKs (Deppermann, Cherpokova et al. 2013). As the incubation time was lengthened, we observed that the level of FGN in Nbeal2-/- MKs became significantly lower than in WT cells, suggesting an altered ability to accumulate FGN. Using FGN multiply-labelled with biotin we looked for evidence of the accumulation of fragments resulting from protein degradation, but these were not detected in Nbeal2-/- MKs. This result indicates that Nbeal2-/- MKs do not show increased degradation of FGN, consistent with reports that MKs do not contain secondary lysosomes (Bentfeld-Barker and Bainton 1982), but rather pass primary lysosomes on to platelets that secrete their contents during activation.

6.4.4 Lack of NBEAL2 results in aberrant secretion of fibrinogen

Examination of FGN secretion revealed that in the absence of NBEAL2, endocytosed FGN is aberrantly externalized by MKs (Fig. 6.11) in a manner akin to MK-synthesized VWF and TSP1. This result indicates that NBEAL2 is required for the retention of MK-synthesized and endocytosed α-granule cargo proteins. It is interesting to note that WT MKs exhibited a moderate release of FGN after extended culture (Fig. 6.11), suggesting that FGN release may be a normal phenomenon that becomes dysregulated in the absence of NBEAL2. It may also be that some surface-bound FGN is released by MKs without being taken into the cell.

6.4.5 Fibrinogen and VWF are abnormally trafficked in Nbeal2-/- megakaryocytes

The trafficking of FGN within MK was examined to gain a better understanding of the role of NBEAL2 in α-granule biogenesis. To do this, the intracellular localization of endocytosed FGN was examined over time relative to an assortment of cell compartment markers. Following its endocytosis by WT MKs, FGN was observed to move to RAB5 early endosomes, RAB7 late

169 endosomes and P-selectin compartments likely indicating maturing α-granules, where FGN was retained (Fig. 6.13A). Initial trafficking of FGN in Nbeal2-/- MKs was similar to WT, but following transient localization with the P-selectin compartment FGN was subsequently transported to RAB11 compartments and then lost from the cells (Fig. 6.13B). Analysis of intracellular VWF in Nbeal2-/- MKs also revealed a patch-like distribution that co-localized with RAB11 compartments (Fig. 6.15). Previously, we have shown that non-permeabilized Nbeal2- /- MKs often show strong peripheral VWF staining (Fig. 5.9). These results suggest that RAB11 is associated with the compartment that releases α-granule proteins in the absence of NBEAL2. Immunogold electron microscopy studies have shown that endocytosed FGN passes through MVBs prior to entry into α-granules (Heijnen, Debili et al. 1998). Here we analyzed in greater detail the trafficking of FGN in MKs, and identified the intracellular compartments involved. Combined with evidence that NBEAL2 co-localizes and interacts with P-selectin, (Fig. 3.10-Fig. 3.12) these results suggest that NBEAL2 is directly involved in the retention of cargo proteins in α-granules.

RAB11 has been implicated in a variety of cellular trafficking pathways, including the post- Golgi secretory pathway and the late recycling endosome pathway, where it regulates the transport of receptors and adhesive proteins and its function is closely related to the exocyst complex (Welz, Wellbourne-Wood et al. 2014). As RAB11 is commonly involved in secretory pathways, it is thus reasonable that RAB11 compartments may also be involved in the release of α-granule proteins. Interestingly, another BEACH domain-containing protein in Drosophila, blue cheese (Bchs), has been reported to function as an antagonist to RAB11 (Khodosh, Augsburger et al. 2006). It may be that NBEAL2 plays a similar role in MKs to inhibit secretion of α-granule cargo via a RAB11-mediated process that is allowed to proceed in its absence.

6.4.6 α-granules externalization is not the direct cause of α-granule protein loss

We have shown that RAB11 may be mediating the aberrant release of α-granule proteins in the absence of NBEAL2. However, one outstanding question is whether in Nbeal2-/- MKs, α- granule surface mobilization contributes to the loss of cargo proteins. Normally, α-granules are mobilized to the cell surface upon cell activation. The secretion of α-granules is not affected by the loss of NBEAL2, as GPS phantom granules contain P-selectin and are secretion competent (Cramer, Vainchenker et al. 1985, Rosa, George et al. 1987). This leads to the fusion of α-granule membrane with the plasma membrane, resulting in the surface

170 expression of α-granule membrane proteins such as P-selectin. By this process, P-selectin can then be detected on the cell surface (Fitch-Tewfik and Flaumenhaft 2013, Kahr, Lo et al. 2013). Using non-permeabilized MKs, our results showed that while VWF is peripheralized in Nbeal2-/- MKs, this is it is not associated with surface mobilization of P-selectin (Fig. 6.17A). This suggests that the secretion of VWF is likely not linked to α-granule surface mobilization. Instead, RAB11 is detected near the cell surface (Fig. 6.17B), which is consistent with our hypothesis that in the absence of NBEAL2, RAB11 mediates VWF secretion.

6.4.7 NBEAL2 is required for the retention of α-granule proteins

The results presented in this study provide novel insights into the function and role of NBEAL2 in α-granule maturation, and the trafficking of α-granule cargo proteins within MKs. Initial endocytosis of FGN into the cell is mediated by integrin αIIbβ3 receptors, and FGN is subsequently transported to RAB5 early endosomes, RAB7 late endosomes then to maturing α-granules containing P-selectin. MK-synthesized α-granule cargo proteins likely follow a similar pathway from the Golgi apparatus to late endosomes or MVBs (Heijnen, Debili et al. 1998) and maturing α-granules, which are stabilized by NBEAL2, consistent with the proposed function of BDCPs as scaffold proteins (Cullinane, Schaffer et al. 2013).

α-granule biogenesis occurs in MKs, but FGN uptake continues in platelets. The trafficking of FGN in platelets is mediated by ARF6 (Huang, Joshi et al. 2016) and VAMP3 (Banerjee, Joshi et al. 2017) where FGN is first transported to RAB4 then to RAB11 compartments. We did not detect an association between FGN and RAB4 in MKs. This may indicate that the trafficking of FGN differs in MKs and platelets, a reasonable expectation given that the latter lack cellular compartments such as the ER and Golgi apparatus (Yadav, Williamson et al. 2017). It should also be noted that unlike the MKs we studied, the platelets used in the endocytosis experiments were loaded with FGN in vivo (Huang, Joshi et al. 2016, Banerjee, Joshi et al. 2017), which may influence the trafficking of the endocytosed FGN.

A clinical manifestation of GPS is the presence of progressive myelofibrosis (Nurden and Nurden 2007, Gunay-Aygun, Zivony-Elboum et al. 2010). α-granules host a diverse spectrum of proteins with various functions. Among these proteins are numerous growth factors and cytokines including TGF-β, FGF, PDGF and PF4 (Maynard, Heijnen et al. 2010). It is likely that the persistent release of these proteins by GPS MKs into the bone marrow will remodel

171 the bone marrow niche through the recruitment of inflammatory cells and fibroblast, thereby potentially leading to the onset of myelofibrosis. Splenomegaly is another hallmark of GPS and is represented by extramedullary hematopoiesis in the spleen (Nurden and Nurden 2007). The release of these growth factors may also contribute to the extramedullary hematopoiesis by promoting the survival, proliferation and growth of the cells in the spleen.

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Chapter 7

Concluding remarks

173

7.1 Functional significance of NBEAL2 molecular and structural properties

The function and role of NBEAL2 was unknown prior to its association with GPS, where the absence of α-granules in the absence of this protein indicated that NBEAL2 plays an essential role in α-granule biogenesis. The first step in examining this function was to explore the molecular and structural properties of NBEAL2. Native PAGE analysis indicated that NBEAL2, a 300 kD protein, is capable of forming 600 kDa or 900 kDa complexes that presumably represent dimers and trimers. As a member of the BDCP family, NBEAL2 is likely to function as a scaffolding protein to facilitate protein-protein interactions (Cullinane, Schaffer et al. 2013), and multimerization may be an aspect of this function by facilitating a range of protein-protein interactions. Further investigation will be needed to explore this aspect of NBEAL2 function.

We have shown that NBEAL2 colocalizes with P-selectin in cells, most likely in the context of α-granules, and that these proteins can also bind in vitro. The details of the interaction of NBEAL2 with P-selectin and α-granules remain to be determined, but our results indicate that NBEAL2 is predominately cytoplasmic and thus likely interacts with the outer surface of α- granules to stabilize their contents. The cytoplasmic distribution of NBEAL2 gives it the potential to participate in a variety of cellular processes, which may account for defects observed in Nbeal2-/- MKs in addition to α-granule deficiency.

The PH-BEACH domain is a hallmark of the BDCP family, and several missense mutations causative for GPS have been identified in the PH-BEACH domain of NBEAL2 (Fig. 1.11). This suggests that the PH-BEACH domain is key to NBEAL2 function, and it will be interesting to examine the molecular consequences (e.g. protein structure, binding of proteins and other molecules) of these mutations. While we were not able to obtain PH-BEACH protein of sufficient quality for structural elucidation, structure modelling using Phyre2 (Kelley, Mezulis et al. 2015) revealed that the PH-BEACH domain mutations, denoted by R2071P, P2100L, H2159Q, H2263Y, S2269L and R2345W, are all positioned in close proximity within what appears to be an active site. We have begun to characterize this domain utilizing a mammalian cell expression system.

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7.2 SEC22B is a novel interacting partner of NBEAL2 that is required for α-granule biogenesis

To gain insights into the functions of NBEAL2, we undertook a search for potential binding partners utilizing AP-MS and megakaryocytic DAMI cells, which do not generate proper α- granules. Among the proteins identified were 14-3-3, FLNA and SEC22B, and we verified binding interaction of NBEAL2 with SEC22B, an integral ER membrane protein. Molecular mapping identified that the 1798-1903aa region of NBEAL2 interacts with SEC22B. Interestingly, two GPS-causing missense mutations, E1833K and R1839C, have been identified in this region. Analysis of NBEAL2 containing these mutations showed loss of interaction with SEC22B, suggesting that NBEAL2-SEC22B interaction is required for α- granule production. To further examine the significance of this interaction we knocked out expression of these proteins in imMKCL cells, a megakaryocytic progenitor line that can be induced to generate α-granules (Nakamura, Takayama et al. 2014), using the CRISPR-Cas9 system. Both SEC22B-KO and NBEAL2-KO imMKCL cells displayed a similar α-granule deficiency, and we also observed that expression of NBEAL2 was greatly decreased in cells lacking expression of SEC22B. These results suggest that SEC22B-NBEAL2 interactions are required for α-granule biogenesis.

SEC22B has been reported to be involved in the trafficking of phagosome contents from the ER (Cebrian, Visentin et al. 2011), and in the transfer of PM lipids through the formation of ER-PM MCS (Petkovic, Jemaiel et al. 2014). These studies provide clues to the potential mechanisms of NBEAL2-SEC22B interaction in α-granule biogenesis. NBEAL2 and SEC22B may form MCS which facilitate the maturation of α-granules by providing structural support and promoting membrane lipid transfer from the ER. It is also possible that NBEAL2 and SEC22B regulate the trafficking of α-granule proteins, although we showed in Chapter 6 that the trafficking of proteins to α-granules is not affected by the loss of NBEAL2. Further studies of NBEAL2-SEC22B interactions would undoubtedly advance our understanding of α-granule biogenesis.

7.3 A model for the role of NBEAL2 in α-granule biogenesis

Based on our data, we propose a model of α-granule cargo protein trafficking in MKs shown in Fig. 6.18. In normal MKs, FGN enters the cell via integrin αIIbβ3-mediated endocytosis. It then passes through RAB5 early endosomes, RAB7 late endosomes or MVBs, and then is retained in the maturing P-selectin α-granules. In the absence of NBEAL2, FGN is

175 endocytosed normally and passes through RAB5, RAB7 and P-selectin compartments, but it then moves to a RAB11 compartment and is released from the cell. MK-synthesized cargo such as VWF is likely trafficked in a manner similar to FGN after its exit from the Golgi, following the classical endocytic pathway (Jovic, Sharma et al. 2010), and in the absence of NBEAL2 this cargo also moves to RAB11-associated vesicles that mediate its release. The NBEAL2 interactome is diverse, and it is likely that NBEAL2 interacts with several proteins to facilitate α-granule stability. Future studies delineating the mechanisms of NBEAL2 interactions with other proteins will provide additional insights into α-granule formation and function.

Figure 7.1. Model of the role of NBEAL2 in α-granule cargo trafficking and maturation. Endocytosed FGN moves from RAB5-positive early endosomes to RAB7 late endosomes, and then to vesicles containing P-selectin. MK-synthesized granule cargo (e.g. VWF, TSP1) follows a convergent route after leaving the Golgi. NBEAL2 is required to stabilize P-selectin vesicles as they mature into α- granules, and in its absence granule cargo is trafficked to RAB11 recycling endosomes and ultimately externalized. This prevents normal α-granule maturation, producing the empty structures observed in Nbeal2-/- mouse and human platelets.

7.4 Implications of NBEAL2 in megakaryocyte and platelet biology

In this thesis, we showed that Nbeal2-/- platelets display impaired activation and aggregation. In conjunction with thrombocytopenia, these defects likely contribute to a bleeding diathesis in Nbeal2-/- mice. Analysis of Nbeal2-/- MKs revealed that NBEAL2 is required for MK maturation, since Nbeal2-/- MKs showed reduced survival, delayed development and reduced proplatelet

176 production leading to thrombocytopenia. GPS is also characterized by progressive myelofibrosis and splenomegaly (Nurden and Nurden 2007, Gunay-Aygun, Zivony-Elboum et al. 2010). In this thesis we showed that NBEAL2 is required for the retention of α-granule cargo proteins, which in its absence are aberrantly released. α-granules contain a host of growth factors, including TGF-β, FGF, PDGF and PF4 (Maynard, Heijnen et al. 2010), and the abnormal release of these factors into the bone marrow and spleen likely contributes to the remodeling of the environment within these organs.

Our results indicate that NBEAL2 is involved in multiple cellular functions in MKs, and likely in platelets as well. Platelets are intimately involved in numerous physiological processes, including hemostasis, immune defense and inflammation, and in pathological conditions such as malignancies and atherothrombosis. Thus understanding NBEAL2 function may aid in the development of new therapeutic strategies to regulate platelet functions in the context of health and disease. Studies aimed at characterizing the molecular interactions of NBEAL2 with binding partners will be of particular relevance.

Prior to this study it had been hypothesized that NBEAL2 was involved in the trafficking of α- granule proteins. Our observation that FGN is transported to P-selectin vesicles similarly in WT and Nbeal2-/- mouse MKs does not support this hypothesis, rather our results indicate that NBEAL2 plays an essential role in the retention of cargo proteins within α-granules. Interestingly, immature α-granules are evidently present in GPS platelets. While these α- granules appear as empty vesicles, indicating the absence of cargo proteins, they are secretion competent as shown by P-selectin surface mobilization (Rosa, George et al. 1987). Similar P-selectin surface mobilization is seen in Nbeal2-/- platelets as we have shown in Chapter 5. In addition to being labelled by P-selectin, the scant presence of VWF and albumin in these vacuoles suggest their identity as immature α-granules (Cramer, Vainchenker et al. 1985, Rosa, George et al. 1987). Based on these observations, it is likely that NBEAL2 is not essential for the generation of these immature α-granules. In contrast to ARC platelets which lack P-selectin (Urban, Li et al. 2012), P-selectin is readily detected in Nbeal2-/- platelets (Kahr, Lo et al. 2013). This suggests the possibility that the VPS16B/33B complex implicated in ARC syndrome may be responsible for generating α-granule structures that NBEAL2 acts to stabilize.

177

Several aspects of α-granule biogenesis still remain to be elucidated, including mechanisms that regulate the trafficking of cargo proteins and generate granule structures. While much progress has been made in studies of proteins specifically involved in α-granule biogenesis identified in studies of inherited deficiencies, it is likely that future progress will involve widening the net to examine the role of proteins with broader cellular functions. These proteins, such as SEC22B, may not be manifested in hereditary disease because loss of expression is lethal. Nevertheless, as our studies have shown, it is possible to study the role of these proteins using cellular models, and presumably lineage-restricted animal models as well.

7.5 Future directions 7.5.1 Molecular and structural properties of NBEAL2

It was predicted from in silico studies that BDCP may cross-interact (Cullinane, Schaffer et al. 2013), and we have shown here that NBEAL2 can form homo-oligomers. Protein multimerization can contribute to stabilization and regulation (Marianayagam, Sunde et al. 2004, Mei, Di Venere et al. 2005), thus it will be of interest to investigate this aspect of NBEAL2 function. If NBEAL2 oligomers have a functional role, then loss of this ability may have consequences, and it is possible that one GPS-associated variant may be deficient in oligomerization, based on Fig. 1.11. Studies of self-binding regions of NBEAL2 via molecular mapping will likely yield useful information, as will experiments where NBEAL2 expression is rescued in NBEAL2-KO imMKCL using variants with and without the capacity to self-interact.

Blue-Native PAGE analysis indicates that NBEAL2 (302 kDa) likely forms homodimers (600 kDa) and trimers (900 kDa). To further define the nature of these multimeric complexes, high resolution analytical techniques such as size exclusion chromatography with inline multi- angle light scattering (SEC-MALS) can be utilized (Sahin and Roberts 2012). Since the possibility remains that the NBEAL2-containing complexes observed may contain other proteins, complexes immunoprecipitated from primary MK lysates can also be analyzed via MS to determine their protein composition. These studies will provide insights into the nature of NBEAL2-containing complexes.

Six GPS-causing missense mutations have been reported in the PH-BEACH region of NBEAL2 (Fig. 1.11), thus studies of this domain will provide key insights into the mechanisms of NBEAL2 function. Future efforts will focus on the expression and purification of the PH-

178

BEACH domain to elucidate its structural properties and potential to interact with other biomolecules. These may include membrane-associated phospholipids (Maffucci and Falasca 2001, Lenoir, Kufareva et al. 2015), which have been reported to interact with PH domains in other proteins. The PH-BEACH domain of LRBA has been reported to not bind phospholipids (Jogl, Shen et al. 2002, Gebauer, Li et al. 2004), while WDFY3 and NSMAF have been reported to be capable of binding phospholipids, suggesting functional heterogeneity among BDCP (Simonsen, Birkeland et al. 2004, Haubert, Gharib et al. 2007).

While we have shown in this thesis that NBEAL2 interacts with P-selectin, it remains possible that their interactions involve other proteins, molecules (e.g. phospholipids) or cell structures (e.g. membranes). One step towards a better understanding of the interaction of NBEAL2 with P-selectin will involve purifying these proteins and examining their direct interaction, as well as the interaction of various subdomains. For example, it should be possible to determine whether NBEAL2 interacts with those parts of P-selectin that extend into the granule lumen or with the cytoplasmic tail portion.

7.5.2 SEC22B as an essential binding partner of NBEAL2 in α-granule biogenesis

In Chapter 4 we determined that SEC22B interacts with NBEAL2, that SEC22B is essential for the formation of α-granules, and that SEC22B-NBEAL2 interactions likely facilitate α- granule maturation. It is possible that SEC22B and NBEAL2 form non-fusogenic MCS, thereby bringing the ER and α-granules in close contact. MCS formation enables the transport of phospholipids from the ER to vesicles, in this case maturing α-granules, where MCS formation may also provide structural support. Future studies will be focused on the characterization of the precise mechanism by which NBEAL2-SEC22B interactions mediate α-granule loading, maturation and stability. For example, initial study aimed to dissect the differences in phospholipid composition of the ghost α-granules from NBEAL2-KO and SEC22B KO imMKCL cells will yield insights into the potential mechanisms of NBEAL2- SEC22B interaction.

It has been reported that CRISPR-mediated gene editing can introduce off-target gene deletions, and it will be important to examine our SEC22B-KO imMKCL cell line for evidence that such deletions may be contributing to the observed α-granule deficiency. This will be done by introducing CRISPR-proof SEC22B into SEC22B-KO imMKCL cells, which if it rescues the α-granule defect will confirm the role of SEC22B in α-granule biogenesis. If the

179 defect is not rescued, the result will be of interest since a screen of the SEC22B-KO cell line is likely to identify one or more other proteins involved in α-granule biogenesis.

7.5.3 NBEAL2 stabilizes α-granules to retain α-granule cargo proteins

We determined that NBEAL2 is likely involved in the stabilization of α-granule cargo, and the question remains as to the mechanisms involved. A promising lead is that NBEAL2 appears to interact with α-granule membranes by binding to P-selectin and/or membrane phospholipids (via the PH-BEACH domain). Future studies will investigate the nature of the interaction of NBEAL2 with α-granules directly, and in the context of interaction with other partners such as SEC22B. It is likely that NBEAL2 performs a scaffolding role that involves interaction with multiple proteins and membranes, and evidence is emerging of an extensive interactome. For example studies in HEK293 cells have identified NBEAL2 interactions with DOCK7, VAC14 and SEC16A (Mayer, Jasztal et al. 2018).

In studies of the BDCP Bchs in Drosophila, it was found that this protein is an antagonist to RAB11-mediated vesicular trafficking events (Khodosh, Augsburger et al. 2006). This raises the possibility that NBEAL2 may similarly act as a RAB11 antagonist to prevent the loss of α- granule proteins. Future studies may delineate the interaction of NBEAL2 with RAB11 in MKs and platelets to characterize their role in α-granule production and stabilization.

7.5.4 Other functions of NBEAL2 in megakaryocytes and platelets

The diverse interactome of NBEAL2 points to multiple cellular functions, as do the many defects seen in Nbeal2-/- mice which suggest that NBEAL2 is involved in MK survival, development and proplatelet production, and in platelet activation and aggregation. While they are as yet poorly understood, some tentative connections can be made. For example, 14-3-3 has been shown to be a central player in mediating signal transduction events involved in platelet activation (Chen, Ruggeri et al. 2018), and FLNA-deficient mice show a macrothrombocytopenia phenotype similar to that seen in Nbeal2-/- mice (Begonja, Pluthero et al. 2015). The loss of NBEAL2 interaction with 14-3-3 and FLNA may contribute to the defects seen in Nbeal2-/- mice. GPS patients show diverse manifestation of symptoms (Nurden and Nurden 2007, Gunay-Aygun, Zivony-Elboum et al. 2010), and it is possible that different NBEAL2 mutations cause varying protein interaction defects that are contributing to

180 this heterogeneity of symptoms. Future studies aimed at characterizing the functional significance of NBEAL2 interactions will provide key insights.

181

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