Characterizing and Quantifying Membrane Order of Polarized Epithelial Cells in Zebrafish Larvae

Home , Epithelial polarity, Lipid raft

Characterizing and quantifying membrane order of polarized epithelial cells in zebrafish larvae

A thesis presented for the degree of

Doctor of Philosophy

By Ahmed Abu-Siniyeh

Faculty of Medicine

Centre for Vascular Research

2014

Originality statement

"I hereby declare that this submission is my own work and that, to the best of my knowledge and belief, it contains no material previously published or written by another person nor material which to a substantial extent has been accepted for the award of any other degree or diploma of the UNSW or any other educational institution, except where due acknowledgment has been made in the thesis. Any contribution made to the research by others, with whom I have worked at UNSW or elsewhere, is explicitly acknowledge in the thesis. I also declare that the intellectual content of this thesis is the product of my own work, except to the extent that assistance from others in the project's design and conception or in style, presentation, and linguistic expression is acknowledged.

Signature......

Name ......

Date ......

‘I hereby grant the University of New South Wales or its agent the right to archive and to make available my thesis or desertion in whole or part in the University libraries in all forms of media, now or hereafter known, subject to the provisions of the Copyright Act 1968. I retain all proprietary rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation.

I also authorize University Microfilms to use the 350 word abstract of my thesis in Dissertation Abstract International.

I have either used no substantial portions of copyright material in my thesis or I have obtained permission to use copyright material; where permission has not been granted I have applied/will apply for a partial restriction of the digital copy of my thesis or dissertation'.

Signed: ------

Ahmed Abu siniyeh

Date: ------

Authenticity statement

‘I certify that the Library deposit digital copy is a direct equivalent of the final officially approved version of my thesis. No emendation of content has occurred, and if there are any minor variations in formatting, they are the result of the conversion to digital format'.

Signed: ------

Ahmed Abu siniyeh

Date: ------

iii

Abstract

The composition and structure of plasma membranes is critical for many cell functions. The plasma membrane of polarized epithelial cells can be divided into two compartments, the apical and basolateral membrane that differ in compositions and function. In particular, it has been proposed that the apical membrane is enriched in lipid rafts. Lipid rafts are defined as small lipid domains that are enriched in cholesterol and sphingolipids. Thus a biophysical property of raft membranes is that they are more ordered than non-raft membranes. The apical-basolateral polarity in polarized epithelial cells is maintained by polarity proteins. Polarity proteins typically localized to either apical or basolateral membranes and organize intracellular trafficking to either membrane compartment. It is currently unknown whether and how membrane organization and polarity networks are linked in polarized epithelial cells. The purpose of the research presented in this PhD thesis was to investigate the relationship membrane order and polarity protein localization.

A better understanding of the cell membrane and its biophysical properties can be achieved by visualizing and analyzing membranes in whole vertebrate organisms rather than imaging cells in tissue culture systems. This is because in vivo, the cellular organization within organs is maintained, which is not just critical for the polarization of epithelial cells but also likely to affect the physical-chemical properties of cell membranes. Here, zebrafish was used as a model organism because its transparency enables high-resolution fluorescence imaging. Zebrafish has become a popular animal model because embryos can be genetically manipulated to study the molecular basis for disease including diseases in which epithelial cell polarization is a key factor.

In this study, the transparency of zebrafish was exploited to study the relationship between membrane order and the localization of polarity proteins in epithelial cells in three different tissues: gut, kidney, and liver. Using the membrane dye Laurdan and multi-photon microscopy, membrane order of polarized epithelial cells in the gut, kidney, and liver were quantified at different development stages. Laurdan incorporates itself into cell membranes parallel to the hydrophobic tails of phospholipids. The probe displays spectral sensitivity to the polarity of its environment, with a ∼50-nm red shift of its emission maximum in polar versus nonpolar environments. This shift in emission profile allows a quantitative assessment of membrane order by calculating a ratiometric measurement of the fluorescence intensity recorded in two spectral channels, known as a generalized polarization (GP) value. A change in membrane order in epithelial cells was observed during development with particularly high membrane order recorded at 6 days post fertilization (dpf) for all three tissues. Apical membranes were significantly more ordered than the basolateral membranes, and basolateral membranes were more ordered than intracellular membranes in gut, kidney, and liver at 3-11 pdf.

Manipulation of 6 dpf larvae with either 7-ketocholesterol (7KC) or cholesterol- methyl-β-cyclodextrin complexes (cholesterol-mβCD) or methyl-β-cyclodextrin

(mβCD) significantly decreased membrane order in the apical, basolateral and intracellular membranes in gut, kidney, and liver epithelial cells. When 4 dpf larvae were treated with 7KC, cholesterol-mβCD or mβCD, the membrane order of apical, basolateral and intracellular membranes was also significantly decreased but recovered to almost normal levels when 4 dpf sterol-manipulated larvae were grown to 6 dpf.

Tissue organization was largely unaffected by the sterol manipulations but apical

targeting of atypical protein kinase C (aPKC) was reduced in sterol-manipulated 4 pdf larvae that also recovered after a further two days of development.

Therefore, a strong correlation between the high membrane order and apical localization of aPKC was observed under conditions where membrane order was acutely decreased and post recovery.

To investigate whether polarity and membrane proteins influence membrane order in polarized epithelial cells, morpholinos (MO) were used to knockdown the expression of the polarity proteins Par3 (par3 MO) and Crb3a (crb3a MO), and the lipid raft proteins, Flotillin-1a (flot-1a MO) and Flotillin-2 (flot-2a MO). A significant decrease in membrane order was found in the apical, basolateral and intracellular membranes in epithelial cells in the gut, kidney, and liver of par3, crb3a, flot-1a, and flot-2a morphants larvae at 4 dpf compared to control. In contrast, a decrease in apical localization of aPKCs was only observed in epithelial cells of par3 and crb3a morphants larvae but not in flot-1a and flot-2a morphants larvae.

In conclusion, the data suggest that membrane order and polarity protein networks are mutually dependent on each other in polarized epithelial cells of the gut, kidney, and liver of intact zebrafish larvae. Decrease in membrane order resulted in a diminished apical localization of the polarity protein aPKC and reduced expression of the polarity proteins Par3 or Crb3a decreased membrane order. This strongly suggests that polarity networks and membrane organization are not controlled by two distinct cellular processes and networks but are part of the same biology. It is possible that the delivery of lipids to apical and basolateral membranes by polarity proteins maintains the differences in order in these membranes and that membrane order is a trafficking

‘signature’ for apical sorting. Hence, the work presented here is only the first step towards a complete picture of the functional relationship between polarity proteins and membrane organization.

The multidisciplinary approach taken here may inspire future studies to better integrate the biology of membranes with the biology of polarized trafficking.

vii

Acknowledgments

I am deeply thankful to Professor Katharina Gaus for offering me the opportunity to engage in PhD and study in her lab. I will not forget the support and encouragement she has shown to me during my study. I have learned a lot from her which I think will be helpful for my career.

I wish to thank Dylan for his technical and scientific advices in microscopy and for helping me in my first and last draft, he was always there when I need his advice. A big thank for Arindam Majumdar for his tremendous help in zebrafish issues as he gave me a big support with my experiments. Thanks also to Thomas Becker for let me use his fish during my PhD project and for all the BRMI friendly staff for their help.

Thanks a lot to my colleagues in the lab, especially Jeremie and Abigail, who have helped me in completing my work successfully. I’d like to thank the staff at the Centre for Vascular Research and at BMIF in particular Alex Macmillan and Michael Carnell.

Special thanks to my best friend Waleed for his encouragement and for his technical advices during my PhD study.

Finally I could never have reached this goal without the fantastic love and emotional support of my family; my mother and my brothers, and a big special thanks to my wife for her support and being patient during all my study.

viii

Table of Contents

CHARACTERIZING AND QUANTIFYING MEMBRANE ORDER OF POLARIZED EPITHELIAL CELLS IN ZEBRAFISH LARVAE ...... 1 ORIGINALITY STATEMENT ...... I COPYRIGHT STATEMENT...... II AUTHENTICITY STATEMENT ...... III ABSTRACT ...... IV ACKNOWLEDGMENTS ...... VIII TABLE OF CONTENTS ...... IX TABLE OF FIGURES ...... XIV ACRONYMS & ABBREVIATIONS ...... XVI CHAPTER 1: INTRODUCTION ...... 1 1.1 ORGANIZATION OF CELL MEMBRANES AND LIPID DOMAINS ...... 1 1.1.1 Plasma membrane components ...... 1 1.1.2 Membrane lipids in cell membrane ...... 1 1.1.3 Cell membrane proteins, cortical cytoskeleton and glycocalyx ...... 5 1.1.4 Membrane phase separation ...... 11 1.1.5 The Lipid Raft hypothesis ...... 14 1.1.6 Definition of Lipid rafts ...... 15 1.1.7 Lipid rafts proteins ...... 17 1.1.8 Lipid rafts and cell functions ...... 20 1.1.9 Lipid rafts and disease ...... 22 1.2 METHODS TO STUDY MEMBRANE ORGANIZATION ...... 23 1.2.1 Biochemical methods ...... 23 1.2.2 Advanced microscopy methods ...... 25 1.2.3 Environmentally sensitive dyes ...... 31 1.3 LIPID RAFTS DEBATES ...... 37 1.4 ORGANIZATION OF POLARIZED EPITHELIAL CELLS ...... 40 1.4.1 Physiological function of polarized epithelial cells ...... 40 1.4.2 Membrane organization in polarized epithelial cells...... 41 1.5 HOW IS THE MEMBRANE ORGANIZATION IN POLARIZED EPITHELIAL CELLS MAINTAINED? ...... 43 1.5.1 Lipid rafts in polarized epithelial cells ...... 43

1.5.2 Membrane trafficking ...... 44 1.5.3 Polarity proteins ...... 46 1.6 ZEBRAFISH AS A MODEL TO STUDY EPITHELIAL CELLS IN VIVO ...... 51 1.6.1 Introduction to zebrafish ...... 51 1.6.2 Structure and function of polarized epithelial cells in zebrafish ...... 53 1.6.3 Zebrafish as a disease model ...... 58 1.7 AIMS OF THIS STUDY ...... 63 CHAPTER 2: MATERIALS AND METHODS ...... 66 2.1 ZEBRAFISH ...... 66 2.1.1 Zebrafish strain and husbandry ...... 66 2.1.2 Set up mating, collecting and sorting zebrafish embryos ...... 66 2.2 LAURDAN MICROSCOPY ...... 67 2.2.1 Laurdan staining and mounting ...... 67 2.2.2 Imaging and image analysis ...... 67 2.2.3 Calibration factor (G-factor) calculation ...... 68 2.2.4 Image analysis and data acquisition ...... 69 2.3 IMMUNOFLUORESCENCE STAINING AND CONFOCAL IMAGING ...... 71 2.3.1 Antibody staining of whole mounted zebrafish ...... 71 2.3.2 Confocal imaging of zebrafish ...... 72 2.3.3 Antibody staining of sectioned zebrafish ...... 73 2.3.3.1 Zebrafish sectioning using a vibratome ...... 73 2.3.3.2 Antibody labelling ...... 73 2.3.4 Confocal imaging ...... 74 2.4 PREPARATION AND QUANTIFICATION OF STEROL-CYCLODEXTRIN COMPLEXES .... 74 2.4.1 Preparation of Sterol-cyclodextrin complexes ...... 74 2.4.2 Quantification of sterol-cyclodextrin complexes ...... 75 2.5 MORPHOLINO KNOCKDOWN EXPERIMENT ...... 76 2.5.1 Morpholino Oligonucleotides (MOs) sequence ...... 76 2.5.2 Microinjection of morpholino into zebrafish embryos ...... 77 2.5.2.1 Micropipette and microinjection chamber plate preparation ...... 77 2.5.2.2 Morpholino loading ...... 77 2.5.2.3 Injection volume calibration ...... 77 2.5.2.4 Zebrafish embryo preparation for microinjection ...... 77 x

2.5.2.5 Morpholino Oligonucleotides preparation ...... 78 2.5.2.6 Zebrafish microinjection ...... 78 2.6 WESTERN BLOT...... 79 2.6.1 Preparation of total lysates ...... 79 2.6.2 Immunoblot ...... 79 2.6.2.1 Preparing cell lysates and electrophoresis ...... 79 2.6.2.2 Polarity and rafts proteins detection ...... 80 2.6.2.3 Stripping the membrane ...... 81 2.7 STATISTICAL ANALYSES ...... 81 CHAPTER 3: MEMBRANE ORDER OF POLARIZED EPITHELIAL CELLS DURING THE DEVELOPMENT OF ZEBRAFISH LARVAE ...... 82 3.1 INTRODUCTION ...... 82 3.2 DEVELOPMENT OF AN IMAGING PROTOCOL FOR MEMBRANE ORDER MEASUREMENT IN ZEBRAFISH LARVAE ...... 87 3.2.1 Laurdan staining of zebrafish larvae ...... 88 3.2.2 Effects of Laurdan on zebrafish development ...... 88 3.2.3 Image analysis and data acquisition ...... 89 3.3 RESULTS OF MEMBRANE ORDER MEASUREMENTS IN DEVELOPING ZEBRAFISH ..... 89 3.3.1 Membrane order of epithelial cells in the gut during development ...... 89 3.3.2 Membrane order of epithelial cells in the kidney during development .... 94 3.3.3 Membrane order of epithelial cells in the liver during development ...... 95 3.4 EFFECT OF PFA FIXATION ON GP VALUES MEASUREMENTS ...... 97 3.5 CONCLUSION AND DISCUSSION ...... 100 CHAPTER 4: EFFECT OF STEROL MANIPULATIONS ON MEMBRANE ORDER AND POLARITY PROTEIN LOCALIZATION IN POLARIZED EPITHELIAL CELLS IN VIVO ...... 103 4.1 INTRODUCTION ...... 103 4.2 STEROL MANIPULATIONS OF INTACT LARVAE ...... 108 4.2.1 Conditions of sterol manipulations of intact larvae...... 108 4.2.2 Effect of sterol manipulations on tissue morphology ...... 109 4.2.3 Membrane order in sterol-manipulated larvae ...... 112 4.2.4 Membrane order of epithelial cells in the gut...... 113 4.2.5 Membrane order of epithelial cells in the kidney ...... 115 4.2.6 Membrane order of epithelia cells in the liver ...... 116

Apical membrane ...... 117 Basolateral membrane ...... 117 Intracellular membrane ...... 118 4.3 RECOVERY OF MEMBRANE ORDER AFTER STEROL MANIPULATIONS ...... 118 4.3.1 Membrane order of epithelial cells in the gut, kidney and liver at 4 dpf 121 4.3.2 Membrane order of gut, kidney and liver epithelial cells post recovery (6 dpf) ...... 123 4.4 EFFECT OF STEROL ON APKC APICAL LOCALIZATION...... 126 4.4.1 aPKC localization in the gut, kidney and liver of sterol-manipulated larvae at 4 dpf ...... 130 4.4.2 aPKC localization in the gut, kidney and liver of post recovery sterol- manipulated larvae at 6 dpf ...... 131 4.5 CONCLUSION AND DISCUSSION ...... 132 CHAPTER 5: REDUCED EXPRESSION OF POLARITY PROTEINS AND MEMBRANE ORDER ...... 137 5.1 INTRODUCTION...... 137 5.2 THE EFFECT OF PAR3 AND CRB3A KNOCKDOWN ON MEMBRANE ORDER ...... 140 5.2.1 Western blot ...... 141 5.2.2 Morpholinos knockdown ...... 141 5.2.3 Laurdan staining and image analysis ...... 143 5.3 PAR3 AND CRB3A KNOCKDOWN RESULTS ...... 145 5.3.1 Membrane order in the gut, kidney, and liver of morphants at 4 dpf ..... 145 5.3.4 Tissue comparison ...... 146 5.4 EFFECT OF PAR3 AND CRB3A KNOCKDOWN ON APKC LOCALIZATION ...... 146 aPKC localization in the gut ...... 148 aPKC localization in the kidney ...... 148 aPKC localization in the liver ...... 149 5.5 CONCLUSION AND DISCUSSION ...... 149 CHAPTER 6: REDUCED EXPRESSION OF FLOTILLIN-1 AND FLOTILLIN-2 ...... 153 6.1 INTRODUCTION...... 153 6.2 THE EFFECT OF FLOTILLIN-1A AND FLOTILLIN-2A KNOCKDOWN ON MEMBRANE ORDER IN ZEBRAFISH LARVAE ...... 154 6.2.2 Western blot ...... 154 6.2.1 Morpholino knockdown ...... 154 6.2.3 Laurdan staining and image analysis ...... 156 xii

6.2.4 Flotillin-1a and Flotillin-2a knockdown results ...... 158 Membrane order in the gut, kidney and liver ...... 158 6.3 EFFECT OF FLOTILLIN-1A AND FLOTILLIN-2A KNOCKDOWN ON APKC LOCALIZATION ...... 159 6.4 CONCLUSION AND DISCUSSION ...... 161 CHAPTER 7: CONCLUSION & DISCUSSION ...... 164 BIBLIOGRAPHY ...... 178 APPENDIX A ...... 207 Publications arising from this study ...... 207

xiii

Table of Figures

Figure 1-1: Structures of the major lipid classes in cell membrane...... 5 Figure 1-2: Schematic representation of the cell membrane...... 10 Figure 1-3: Schematic representation of liquid ordered (lo) and liquid disordered (ld) phases in cell membrane...... 12 Figure 1-4: Schematic representation of Flotillin-1 and Flotillin-2 in plasma membrane…… 19 Figure 1-5: Schematic of the emission shift in polar and non-polar environments……………. 32 Figure 1-6: Schematic representation of localization of Par & Crumbs complexes………….. 48 Figure 2-1: Generating GP images and pseudo-colored images for data acquisition……….… 71 Figure 3-1: Wild type-AB zebrafish and mutant strain Casper...... 87 Figure 3-2: Laurdan staining has no effect on zebrafish development...... 89 Figure 3-3: GP image of zebrafish gut...... 90 Figure 3-4: GP images of zebrafish gut, kidney, and liver in 3-11 dpf development stages...... 91 Figure 3-5: Comparison of GP values of apical, basolateral and intracellular membranes in epithelial cells of the gut, kidney, and liver in 3-11 dpf zebrafish larvae...... 92 Figure 3-6: GP image of zebrafish kidney...... 94 Figure 3-7: GP image of zebrafish liver...... 96 Figure 3-8: Changes in GP values ...... 97 Figure 3-9: Comparison of GP values of the apical, basolateral, and intracellular membranes between live and fixed larvae...... 98 Figure 3-10: GP values of apical, basolateral, and intracellular membranes at fixed 6 dpf larvae...... 99 Figure 4-1: Tissue morphology of gut, kidney, and liver in sterol treated 6 dpf zebrafish larvae...... 1109 & 110 Figure 4-2: Membrane order in sterol treated 6 dpf zebrafish larvae...... 113 Figure 4-3: Illustration of sterol manipulation experiment...... 119 Figure 4-4: Membrane order of sterol-manipulated larvae at 4 pdf and after a recovery period of two additional days (6 pdf)...... 120 Figure 4-5: Recovery of membrane order after sterol modification………………………..… 126 Figure 4-6: aPKC apical localization in larvae treated with 7KC, cholesterol-mβCD, and mβCD at 4 dpf and after a 2-day recovery (6 dpf)...... 129 Figure 5-1: Immunoblot of knockdown Par3 and, Crb3a proteins...... 142

xiv

Figure 5-2: Phenotype of larvae injected with par3 MO and crb3a MO Morpholinos...... 143 Figure 5- 3: Membrane order in par3 and crb3a morphants larvae...... 144 Figure 5-4: aPKC localization in the epithelial cells of the gut, kidney, and liver in par3 and crb3a morphants larvae...... 147 Figure 6-1: Immunoblot of knockdown Flot-1a and Flot-2a proteins...... 155 Figure 6-2: Phenotype of injected larvae with flot-1a MO and flot-2a MO Morpholinos…….156 Figure 6-3: Membrane order in flot-1a and flot-2a morphants larvae...... 157 Figure 6-4: aPKC localization in epithelial cells of the gut, kidney, and liver in flot-1a and flot- 2a morphants larvae...... 160 Figure 7-1: Correlation between membrane order and aPKC localization………….………...173

Acronyms & Abbreviations

7KC 7-ketocholesterol AD Alzheimer Disease ADPLD Autosomal dominant polycystic liver disease AFM Atomic force microscopy AJ Adherens junction ALD Alcoholic liver disease ALR Augmenter of Liver Regeneration AP-1 Activator protein 1 aPKC Atypical protein kinase C APP Amyloid precursor protein ASIP Isotype-speciﬁc interacting protein Aβ Amyloid-β-peptide Baz Bazooka BSA Bovine serum Albumin C-laurdan 6-dodecanoyl-2-[N-methyl-N-(carboxymethyl) amino-naphthalene COP Coatomer protein complex Crb Crumbs protein CTxB Cholera Toxin-B DALDA [D-Arg2,Lys4]-dermorphin-(1–4)-amide DIGs Detergent-insoluble glycolipid-enriched complexes Dlg Disc-large DMSO Dimethyl Sulfoxide DOPC 1, 2-Dioleoyl-sn-glycero-3-phosphocholine dpf days post fertilization DPH 1, 6-diphenyl-1, 3, 5-hexatriene DPPC 1, 2-Dipalmitoyl-sn-glycero-3-phosphocholine DRMs Detergent-resistant membranes DSPC 1,2-distearoyl-sn-glycero-3-phosphocholine ECL Enhanced Chemiluminescence ECM Extracellular matrix ER Endoplasmic reticulum ESR Electron spin resonance

xvi

EtOH Ethanol FCS Fluorescence correlation spectroscopy Fgf Fibroblast growth factor FLIM Fluorescence-lifetime imaging microscopy Flot-1 Flotillin-1 protein Flot-2 Flotillin-2 protein FPV Fowl plague virus FRET Fluorescence resonance energy transfer GalNAc N-acetyl galactosamine GlcNAc N-acetyl glucosamine GLUT1 Glucose transporter 1 GP Generalized polarization GPI Glycosylphosphatidylinositol GPMVs Giant Plasma-Membrane Vesicles GSLs Glycosphingolipids GUVs Giant unilamellar vesicles hpf hours post fertilization IBD Inflammatory bowel disease ICM Intermediate cell mass Klf4 Krüppel-like factor 4 LAT Linker for Activation of T cells Laurdan 6-dodecanoyl-2-dimethylaminonaphthalene Lck Lymphocyte-specific protein tyrosine kinase ld Liquid disorder phase LDL Low density lipoprotein LgI Lethal (2) giant larvae lo Liquid-ordered phase LRP1 Receptor-related protein 1 MAL Myelin and lymphocyte protein MDCK Madin-Darby Canine Kidney cell line. MHC Class II antigen presentation MOR μ-opioid receptor mβCD Methyl-β-cyclodextrin NAFLD Nonalcoholic fatty liver disease

xvii

NGS Normal goat serum Par Partition defective protein PATJ PALS1-associated TJ protein PBI Posterior blood land PBS Phosphate buffer saline PC Phosphatidylcholines PD Parkinson's disease PDZ Postsynaptic-density-95 PE Phosphatidylethanolamines PI(3,4,5)P3/PIP3 Phosphatidylinositol (3,4,5)-triphosphate PI(4,5)P2/PIP2 Phosphatidylinositol 4,5-bisphosphate PKD Polycystic kidney disease PLD Polycystic liver disease PMT Photomultiplier tubes POPC 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine PTEN Phosphatase and tensin homolog PTU 1-phenyl 2-thiourea RGCs Retinal ganglion cells Scrb Scribble protein SD Standard Deviations SDS Sodium dodecyl sulphate SJ Septate junction SNARE Soluble NSF attachment protein receptor SOPC 1-stearoyl-2-oleoyl-sn-glycero-3-phosphocholine SPFH Stomatins, Prohibitins, Flotillins, H.K/C SPT Single-particle tracking TCZs Transient confinement zones TGN Trans-Golgi network TJ Tight junction Tm Tunicamycin TNBS Hapten 2,4,6-trinitrobenzene sulfonic acid TNF Tumor necrosis factor UPR Unfolded protein response VSV Vesicular stomatitis virus

xviii

WGA Wheat Germ Agglutinin ZA Zonula adherens ZO-1 Zona-occludens -1

xix

Chapter 1: Introduction

1.1 Organization of cell membranes and lipid domains

1.1.1 Plasma membrane components

As one of the most prominent cell components, the plasma membrane provides a barrier interface between the cell external environment and cytoplasm. It has a fundamental role in a variety of cellular functions such as nutrient transportation, enzymatic catalysis and mediates cell-to-cell communication. In addition, the cell membrane has a vital role in cell signaling by responding to external signals and transmitting them into the cell (Fielding 2006).

The plasma membrane is composed of a lipid bilayer of mainly amphiphilic phospholipids, glycolipids and cholesterol, and different types of membrane-embedded proteins that perform defined functions in the cell membrane. Phospholipids have a hydrophobic "tail" which make the interior of the bilayer inaccessible to the surrounding water while the hydrophilic "head" is the exterior surfaces of the bilayer (Lingwood and

Simons 2010).

1.1.2 Membrane lipids in cell membrane

Phospholipids are a key component of the cell membrane and the most abundant lipids in mammalian cells. Phospholipid fatty acids chains typically contain between 16 to 20 carbon atoms and these fatty acids are either saturated or unsaturated.

Phospholipids in the cell membrane usually exist in liquid-ordered phase under physiological conditions.

Phospholipids play a crucial role in compartmentalizing cells and can also be signaling molecules, help anchor proteins to membranes and can act as energy source.

Phosphatidylcholines (PC), phosphatidylethanolamines (PE) and phosphatidylinositol

(PI) are examples of types of phospholipids that exist in the cell membrane (Von Heijne and Rees 2008; Campbell and Reece 2005; Nelson and Cox 2004).

Glycolipids are lipids that are linked to a carbohydrate. The glycolipid backbone is composed of short, branched chains with less than 15 sugar units. The main difference between glycolipids and sphingomyelin is that glycolipids are linked to the primary hydroxyl group of the sphingosine backbone, while sphingomyelin consists of a ceramide unit and a phosphorylcholine moiety attached to position one .Glycolipids have structured asymmetry in the sugar residues on the external surface of the membrane. In addition, complex structures of glycolipids may include a branched chain of up to seven sugar residues. Glycolipids are markers for cellular recognition due to their location on the outer surface of the cell membrane and provide energy as well as performing other essential functions within the cell. There are different types of glycolipids such as glyceroglycolipids, glycosphingolipids, in addition to cerebrosides and gangliosides, which are the most complex glycolipids and contain negatively charged oligosaccharides with one or more sialic acid residues (Von Heijne and Rees

2008; Campbell and Reece 2005; Nelson and Cox 2004) (Figure 1-1).

Cholesterol is composed of four linked hydrocarbon rings and this ring system is more rigid than other membrane lipids (Figure 1-1). Cholesterol is crucial in forming and maintaining the stability of the cell membrane; depending on the physiological temperatures, cholesterol can control membrane fluidity.

Cholesterol reduces membrane fluidity via the interaction of its hydroxyl group with the polar head groups of phospholipids and sphingolipids. Cholesterol is also involved in cell signaling and it is essential for the function and structure of caveolae and clathrin- coated pits (Von Heijne and Rees 2008). Bretscher and Munro (1993) suggested that the cholesterol content could increase the thickness of the lipid bilayer.

Sphingolipids are a complex range of lipids defined by their fatty acids linked to a long-chain base or sphingoid, which is a set of amino-alcohol backbones, via amide bonds. Sphingolipids have diversity of structures that play important roles in cell function such as signal transmission, cell recognition, and formation of lipid rafts. For example, a sphingolipid with one hydrogen atom only is a ceramide while other R groups such as phosphocholine, produce a sphingomyelin, and different carbohydrate monomers or dimers, produce cerebrosides and globosides, respectively that are together known as glycosphingolipids (Gault et al. 2010) (Figure 1-1).

Lipid bilayers are composed of phospholipids, which have a hydrophilic head and two hydrophobic tails each. These phospholipids arrange themselves as a bilayer with all of their tails pointing toward the center of the layer, when they are exposed to water.

These lipids are arranged heterogeneously within the leaflets themselves and lead to lipid asymmetry in the bilayer (Klausner et al. 1980). Exoplasmic leaflets are rich in phosphatidylcholine, sphingolipids, glycosphingolipids, and sphingomyelin.

Phosphatidylethanolamine and phosphatidylserine are mainly localized to the inner leaflet (Simons and Meers 1988). It is hypothesized that cholesterol is found in both leaflets, but is more abundant in exoplasmic leaflet. This causes lipid asymmetry that depends on the cell type and the activation status of the cell (Binder et al. 2003).

On the other hand, others have suggested that cholesterol distributed more at the cytosolic leaflet (the inner leaflet) of the plasma membrane and the endocytic recycling compartment of a CHO cell line, which is thought to be responsible for membrane domain formation. In this study, the authors observed the vulnerability of dehydroergosterol (DHE) and cholestatrienol (CTL) in CHO cells to quenching by 2, 4,

6-trinitrobenzene sulfonic acid (TNBS) and by slowly flipping lipid-based quenchers.

This study may resolve some of a few doubts about the cholesterol distribution between the inner and outer leaflet, which came from the shortage of appropriate approaches to explore cholesterol distribution in living cells (Mondal et al. 2009). In addition, the cholesterol flip-flop rate in biological membranes (Steck et al. 2002) may make the interpretation of the techniques used to analyzed cholesterol distribution challenging.

Cholesterol may keep flipping during the assay to fill the gaps that are caused due to lipid flipping, lipid metabolism and lipid deformation. Further investigations are required to understand the asymmetric distribution of cholesterol between the two leaflets in the plasma membrane and the endocytic membrane (Mondal et al. 2009). To maintain bilayer asymmetry, enzymes such as flippases and floppase are responsible for arranging lipids across the bilayer in a proper way (Simons and Meers 1988).

Figure 1-1: Structures of the major lipid classes in the cell membrane. A) Cholesterol is based on a four linked hydrocarbon rings forming the bulky steroid structure. B) Glycerophospholipids are based on diacylglycerol and normally carry acyl chains of 16–18 carbon atoms, one of which has a cis double bond. The head group (R) could be neutral, (serine or inositol) to yield a net acidic charge, or basic (ethanolamine or choline) to yield a neutral, or zwitterionic lipid. Glycolipids are typically linked to the primary hydroxyl group of the sphingosine backbone. C) Sphingolipids are based on a ceramide, and its fatty acids are linked to sphingoid, which is typically saturated, or a long-chain base varying from 16-26 carbons, via amide bonds. The head group is either choline (sphingomyelin), or phosphate replaced with glucose (glycosphingolipids), which is further expanded to make a wide range of glycolipids. (Munro 2003).

1.1.3 Cell membrane proteins, cortical cytoskeleton and glycocalyx

Membrane proteins

The cell membrane contains many proteins that have different functions such as receptors and channels. These proteins can be divided to three classes (transmembrane, peripheral and lipid anchored proteins) depending on how they interact with the lipid bilayer.

Integral proteins or transmembrane proteins have amphipathic features, similar to phospholipids. They interact with the inner lipid bilayer through their hydrophobic

regions, while the hydrophilic regions on the other face the surrounding aqueous environment. A number of membrane proteins located in the cytosol are linked to the inner leaflet of the lipid bilayer, either through covalent interaction with the lipid chain or by the amphipathic α-helix located on the protein surface. Iron channels, proton pumps, G protein-coupled receptors are examples of these proteins (Alberts et al. 2002).

One example of transmembrane proteins is the Linker for Activation of T cells (LAT) which localized in raft microdomains and function as a docking site for SH2 domain- containing proteins (Horejsí 2004) (Figure 1-2).

Peripheral membrane proteins are either attached with the peripheral regions of lipid bilayer or linked to integral membrane proteins by non-covalent interactions.

Peripheral proteins interact with the bilayer transiently, because they can dissociate from the membrane to carry out their functions in the cytoplasm. These proteins can be isolated from the membrane by simple extraction procedures that leave the lipid bilayer intact (Alberts et al. 2002) (Figure 1-2).

Lipid-anchored proteins are proteins that are post-translationally modified with acyl groups and they are not directly in contact with the membrane bilayer; instead, they covalently attached via particular oligosaccharides to single or multiple lipids that exist on the outer surface of the membrane. GI-anchored proteins is an example of these proteins (Alberts et al. 2002) (Figure 1-2).

The Glycocalyx

Membrane proteins are often linked with carbohydrates, i.e are glycosylated, which cover the outer surface of the membrane. The surface of the cell coated with carbohydrates is known as the glycocalyx. Carbohydrates are typically linked to 6

asparagine side chains by N-glycosidic linkages and to serine and threonine side chains by O-glycosidic linkages. The associated carbohydrate is commonly either the N-acetyl glucosamine (GlcNAc) or the N-acetyl galactosamine (GalNAc). Proteins linked to an oligosaccharide are known as glycoproteins while those with broader polysaccharide moieties are known as proteoglycans. The major function of the glycocalyx is to protect the cell surface from the outside environment and to prevent unfavorable protein-protein interactions. It can also act as surface markers for different cell recognition processes because of the abundance of oligosaccharides and their exposed position on the cell surface (Alberts et al. 2002).

Cytoskeleton

The cytoskeleton is a fundamental structure of the cell as it determines cell shape and the overall organization of the cytoplasm. It is also responsible for cell movement.

The cytoskeleton is composed of three types of protein filaments: actin filaments, intermediate filaments, and microtubules. These proteins are linked to each other and are attached to the cell membrane through different types of proteins.

Microfilaments (actin filaments) are composed of thin fibers of polymers of actin subunits that attach to specific regions on the plasma membrane. These filaments are responsible for contacting neighboring cells and tissue components as well as attaching the cytoskeleton to the adhesion sites on the cells. Large bundles of actin filaments known as stress fibers can attach to distinct regions called focal adhesions via interaction with integrins. Actin filaments also serve as paths for myosin molecules to bind and move along the microfilament. In intestinal epithelial cells, actin filaments are responsible for forming finger-like extensions of the cell membrane called microvilli.

These microvilli form a brush border, which is a layer on the apical surface that is responsible for absorption. These actin bundles of microvilli attach to the plasma membrane by the calcium-binding protein calmodulin together with myosin I. To stabilize the microvilli, the actin bundles affix to a spectrin-rich region of the actin cortex called the terminal web. The linkage of the actin cytoskeleton with the plasma membrane is thus essential to cell structure and function (Cooper 2000).

The relation between actin filaments and membrane ordered domains has been explored over the last decade, as establishing and maintaining membrane domains often require the actin cytoskeleton. In one study, FRET was used to investigate the role of the actin cytoskeleton in the co-clustering of membrane raft-associated fluorescent proteins and fluorescent proteins targeted to the non-raft membrane domains of T cells.

They observed a particular co-clustering of raft-associated donor and acceptor probes when the actin cytoskeleton was disrupted by latrunculin (Lat B) (Chichili and Rodgers

2007). On the other hand, the co-clustering of the raft-associated fluorescent proteins increased more than that of the non-raft probes after treatment with jasplakinolide, which increases actin polymerization. Similar effects on protein co-clustering and Lck regulation were observed in cells where cholesterol was sequestered using filipin. This study showed the central role of cholesterol and the actin cytoskeleton in stimulating co- clustering of raft-associated proteins and how this plays a significant part in regulating raft-associated signaling proteins such as Lck (Chichili and Rodgers 2007). In another work, a so-called Ising model, which comprises distinct variables that represent magnetic dipole moments of atomic spins that allow the identification of phase transitions, was used to explore the heterogeneity of a biological plasma

membrane combined with its cortical cytoskeleton. In this study, the thermodynamic basis of heterogeneity in living cell membranes was clarified. The cortical cytoskeleton was shown to play a fundamental part in providing operatively long-term forces between membrane proteins that regulate their organization and dynamics through inducing critical fluctuations in the plasma membrane. This implies that membrane heterogeneity that is modulated by cortical actin could have direct involvements in many cell functions (Machta et al. 2011).

Gowrishankar et al. 2012 hypothesized that lipid-anchored proteins that interact with dynamic actin must display irregular concentration fluctuations, and cell membrane proteins able to connect directly to actin can form nano-clusters. Therefore, by using FCS and TIRF microscopy, they showed that actin filaments and membrane proteins interact and bind to each other to form protein nano-clusters within the cell membrane. This provides new evidence that actin filaments have a role in establishing ordered membrane domains, revealing an active process for the molecular organization of the plasma membrane and its spatiotemporal arrangement. Later, the relationship between actin filaments and plasma membrane order was also confirmed by using laurdan and di-4-ANEPPDHQ in vivo. It was observed that when actin filaments underneath the plasma membrane correlated with ordered domains. This led to the suggestion that the attachment of actin filaments to the plasma membrane has an important role in the formation of ordered membrane domains in vivo (Dinic et al.

2013).

Intermediate filaments have a diameter of approximately 10 nanometers and are more robust than actin filaments. They maintain the shape of the cell, are attached to

organelles, and fix the inner dimensions of the cell. They have significant roles in some cell-to-cell and cell-to-matrix connections. Different kinds of intermediate filaments form from different compounds such as keratin, lamin, and vimentins (Cooper 2000).

Microtubules are highly dynamic hollow cylinders that usually contain 13 protofilaments, which are polymers of alpha and beta tubulin. They are organized by the centrosome. Microtubules are responsible for intracellular transport, maintaining the flagella and cilia, and have an important functional role in mitotic spindle (Cooper

2000).

Figure 1- 2: Schematic representation of the cell membrane. The plasma membrane contains different lipids and proteins that are associated via non-covalent linkage. Phospholipids (shown in blue) are the main and most abundant component of the plasma membrane. In addition, the plasma membrane contains glycolipids (shown in green), which are mainly localized in the outer leaflet with their carbohydrates chain on the cell surface. Cholesterol (shown in red) incorporates itself between phospholipids and is considered one of the main components of lipid rafts in addition to sphingolipids (shown in brown). Different kinds of proteins also exist in the plasma membrane including integral proteins such as LAT (shown in dark brown), which insert themselves through the plasma membrane and interact with inner lipid bilayer through their hydrophobic regions, or linked to carbohydrate chains such as glycoproteins (shown in yellow). Peripheral proteins are either attached to peripheral regions (shown in red) or linked to integral proteins. Lipid-anchored proteins such as GPI-anchored protein (shown in blue) are linked covalently to the lipid or fatty acid chains.

1.1.4 Membrane phase separation

The lipid bilayer is mainly held together by non-covalent forces (hydrophobic interaction) so that lipids can move laterally within the bilayer in a rapid and natural process called "lateral diffusion.” Lipid molecules can also move from one layer to the other in a slow process called "transverse diffusion" or flip-flop movement (Binder et al.

2003). Membrane proteins are also able to move laterally within the cell membrane but their movement is much slower than that of lipids. However, they are not able to move in a transverse manner between the bilayer leaflets (Eeman and Deleu 2010).

At physiological temperatures, the cell membrane usually exists in the liquid state since amphiphilic lipids form a fluid lamellar bilayer in an aqueous environment. The fluidity of the cell membrane is essential for cellular functions to occur. Fatty acid and cholesterol play a crucial role in phase segregation and membrane fluidity. Fatty acid chains with unsaturated cis or trans double bonds increase membrane fluidity; in other words, the fluidity of the membrane may be affected by the length of fatty acid chain and the number of double bonds (Alberts et al. 2009).

Cholesterol is considered the key controller of membrane fluidity due to its ability to merge itself inside the membrane bilayer. The rigid steroid rings and the hydrocarbon chain of cholesterol prefer to interact with saturated lipids rather than unsaturated lipids, while its hydroxyl group interacts with the polar head groups of the neighboring phospholipids, leading to a decrease in membrane fluidity (Simons and Vaz 2004).

Two lipid phases exist in the bilayer membrane, the liquid-ordered phase (lo) and liquid-disordered phase (ld) (Figure 1-3). Janiak and co-authors hypothesized that in the liquid-ordered phase, lipids are arranged on a two-dimensional triangular lattice in the 11

plane of the membrane (Janiak et al. 1979). In phase segregation, the membrane bilayer converts from a highly liquid-ordered phase to a disordered liquid phase and vice versa.

Interestingly, cholesterol and phospholipids in the bilayer have the ability to form a liquid-ordered (lo) phase that can coexist with a liquid-disordered phase (ld), which has a lower concentration of cholesterol. Therefore, cholesterol-dependent domains enriched in sphingolipids and saturated phospholipids can form a liquid-ordered (lo) phase, which is surrounded by liquid-disordered (ld) medium. Lipid phase segregations are mainly influenced by temperature, the chemical structure of the lipid and the presence of cholesterol among the phospholipids (Ipsen et al. 1989; Eeman and Deleu

2010).

Figure 1-3: Schematic representation of liquid ordered (lo) and liquid disordered (ld) phases in cell membrane. Liquid ordered phases contain predominantly saturated phospholipids, sphingolipids, a higher concentration of cholesterol, and a higher lipid packing density compared to liquid disordered phases that contain mainly unsaturated phospholipids.

Lipid phase segregation in membranes with cholesterol and different types of phospholipids have been characterized in different model systems using mixtures of synthetic lipids in monolayers, supported bilayers, and giant vesicles (Hagen and

McConnell 1997; Keller et al. 2000; Dietrich et al. 2001; Veatch and Keller 2002). In these studies, the outcomes indicate that the presence of cholesterol in lipid mixtures produces liquid-liquid phase separation into a (lo) liquid-ordered and a (ld) liquid- disordered phase (McConnell and Vrljic 2003; Simons and Vaz 2004).

A number of scientists have used Giant Plasma-Membrane Vesicles (GPMVs), which are cell-derived liposomes that contain integral and peripheral membrane proteins and a number of different lipids. These giant vesicles showed ld-lo phase separation, similar to that observed in model systems but only under certain conditions. This phase separation was found to separate known protein and lipid markers of lipid rafts, providing a link between ld-lo phase separation in model systems and the lipid raft hypothesis in cellular plasma membranes (Baumgart et al. 2007; Sengupta et al. 2009).

Despite this, using GPMVs reinforced the raft hypothesis as it delivers a useful platform for exploring the compositions and properties of the phases, however, GPMVs still have some limitations as phase separation in GPMVs is restricted to particular environments.

GPMVs were also used as a model to explore the physical basis of the transmembrane linker for activation of T cells (LAT) partitioning into lipid rafts. This study showed that palmitoylation has a pivotal role in regulating raft phase association for integral raft proteins. The outcome of this study proposed that palmitoylation has a role as a dynamic raft targeting mechanism for transmembrane proteins (Levental et al.

2010). In another study, a strong linkage between the miscibility of GPMV and the

relative compositions and order of the coexisting phases was described. In addition, domains with a range of variable stabilities, orders, and arrangements were observed to be induced by quite small differences in isolation conditions. The results of this study showed that domains of different features can be created in a membrane environment whose complexity is reflective of biological membranes (Levental et al. 2011).

Model membranes, which are composed of bilayer mixtures of lipids, are employed by many researchers as chemically simple models for biological membranes. On the other hand, cellular membranes comprise a huge number of lipids and proteins. In term of components, model membranes do not resemble biological membranes, nevertheless scientists implement their studies on model membranes because they display a well- recognized phase segregations (Feigenson 2007).

1.1.5 The Lipid Raft hypothesis

After years of studying membrane microdomains that are enriched with cholesterol and sphingolipids, Karnovsky and his colleagues proposed that there were several phases in the membrane lipid environment by observing heterogeneity in the fluorescence decay of the probe 1, 6-diphenyl-1, 3, 5-hexatriene (DPH), which localizes in the hydrophobic acyl region of the membrane. They also examined the functional effect of altering the membrane structure by the addition of specific fatty acids and the depletion of cholesterol in the cell membrane (Karnovsky et al. 1982).

Later, the hypothesis of lipid rafts was proposed by Simons and his colleagues. The hypothesis declared that the phase behavior of different lipid species is exploited to create lateral heterogeneity in the plasma membrane.

Moreover, they hypothesized that in plasma membrane, unsaturated phospholipids form a liquid-disordered phase that coexists with liquid ordered phase, which is formed from saturated phospholipids and sphingolipids in addition to cholesterol (Simons and Ikonen

1997; Simons and Meers 1988).

Simons and other scientists implemented the concept of membrane microdomains as a foundation for explaining the transport of glycosylphosphatidylinositol GPI- anchored proteins to the cell membrane via the trans Golgi network. The authors revealed the significance of these microdomains as a basis of lipid sorting in vivo, particularly when they noted the enrichment of glycosphingolipids (GSLs) at the apical surface of polarized epithelial cells. These microdomains were named lipid rafts, and the raft theory was proposed as an explanation that lateral segregation in the cell membrane could be supported through the self-associations of sphingolipids and cholesterol. This hypothesis unlocked the door for scientists to investigate new ways of resolving cell membrane enigmas through studying their cell components, structure, functions, and dynamics (Simons and Ikonen 1997; Brown and London 1998).

1.1.6 Definition of Lipid rafts

"Lipid rafts are small (10-200 nm), heterogeneous, highly dynamic, sterol- and sphingolipids-enriched domains that compartmentalize cellular processes. Small rafts can sometimes be stabilized to form larger platforms through protein-protein and protein-lipid interactions." This definition of lipid rafts was developed at the 2006

Keystone Symposium of Lipid Rafts and Cell Function (Pike 2006). Others called these microdomains detergent-insoluble glycolipid-enriched complexes (DIGs), or detergent-

resistant membranes (DRMs) because of the detergent resistance of their constituents

(Fivaz 1999).

Cholesterol and sphingolipids are localized in the exoplasmic and inner cytoplasmic leaflet of the lipid bilayer while sphingolipids are highly concentrated in the exoplasmic leaflet. The heterogeneity of lipid rafts may be due to the differences in protein and lipid composition, which, in turn, is based on the cell type and their activation status. In model membranes, hydrocarbon chains in liquid-ordered phase, such as sphingolipids and phospholipids, are tightly packed and more saturated than in the surrounding bilayer (Brown and London 1998).

Cholesterol has a pivotal role in keeping rafts together as it has the ability to pack in between the lipids in rafts, and fill up the spaces between sphingolipids (Fantini et al.

2004; Korade and Kenworthy 2009). Sphingolipids often interact with cholesterol and enhance the interlocking of the two bilayer leaflets due to the saturation of the elongated hydrocarbon chains (Ramstedt and Slotte 2006). Studies show that interaction with cholesterol forces neighboring hydrocarbon chains into more extended conformations, which affects the thickness of the lipid rafts and the surrounding membrane and promote further segregation between the two phases through hydrophobic mismatches

(García-Sáez et al. 2007; Pike 2009).

Lingwood and Simons (2010) suggested that there are three model states for membrane rafts in living cells, starting with the resting state, in which rafts exist as nano-scale assemblies; then rafts form a platform that leads to its stabilization, and finally, merge to form a coalesced phase. Proteins of lipid rafts are thought to have a vital role in nano-scale assemblies, and to promote the formation of stabilized rafts in

their functional state (Lingwood and Simons 2010). Others hypothesized that rafts merging may establish a platform for new protein-protein interactions that exist on different rafts in the resting state. There is also an indication that differences among lipid raft components can influence the structure, stability, and function of the raft (Pike

2004).

1.1.7 Lipid rafts proteins

It has been proposed that lipid rafts produce a lateral lipid heterogeneity in the cell membrane, leading to a lateral protein heterogeneity. This lateral lipid/protein heterogeneity is thought to be the base for lipid raft function in cells. Furthermore, proteins segregation into microdomains may attract other regulatory proteins to lipid rafts (Bandorowicz-Pikuła 2000).

There are various types of proteins that are thought to associate with lipid rafts either temporarily or permanently. GLUT1, LDL receptor-related protein 1 (LRP1), Lck, and TNF receptor 1 are examples of proteins that temporarily associate with rafts, while caveolins, Flotillins/Reggies, stomatin, LAT, MAL and prominin are examples of permanently associated proteins. Different mechanisms have been hypothesized as to how proteins associate with lipid rafts. These mechanisms include: (1) hydrophobic membrane-spanning sequences or transmembrane domains, (2) hydrophobic tails such as (A) GPI anchors, (B) N-myristoylation or (C) S-palmitoylation, and (3) protein- protein and protein-lipid interactions (Lucero and Robbins 2004).

Membrane associated-proteins, particularly rafts proteins, play an important role in cell functions, and studying the way and how they interact with other membrane components, such as lipids, will clarify more of their contributions to cell functions. 17

One example of putative rafts protein is Flotillin-1 and Flotillin-2. These proteins are assumed to be a key part in different cellular processes such signaling, endocytosis and interactions with the cytoskeleton. Those two proteins are expressed ubiquitously in all mammalian tissues, but there is yet more to understand the molecular details of their various roles. Many studies have been implemented to explore how Flotillins might function in different cellular processes, and it has been proposed that Flotillins operate in different ways in the same process (Nichols and Otto 2011).

The Flotillin (known as the Reggie) protein family consists of highly conserved proteins and is composed of two highly homologous isoforms, Flotillin-1 (Reggie-2) and Flotillin-2 (Reggie-1), which are likely to be expressed ubiquitously (Lang et al.

1998). Flotillins belong to the SPFH (Stomatins, Prohibitins, Flotillins, H.K/C) protein superfamily, and the two isoforms are closely related, Flotillin-1 (~ 47 kDa) and

Flotillin-2 (42 kDa). Flotillins are abundant in the plasma membrane and are considered to be lipid raft markers, where they are linked to the internal leaflet through hydrophobic amino acid extends; they also attach to acyl groups of palmitic and myristic acids while Flotillin-2 is irreversibly myristoylated and multiply palmitoylated

(Browman et al. 2007; Langhorst et al. 2005a) (Figure 1-4). Flotillin-1/Reggie-2 and

Flotillin-2/Reggie-1 are a homo- and hetero-tetramers at the plasma membrane, and it is thought that Flotillin-2 is required for the stabilization of Flotillin-1 (Solis et al. 2007).

Flotillins/Reggies have significant roles in different cell processes in a variety of cell types and tissues, particularly in signal transduction since Flotillins/Reggies have the potential to recruit signaling molecules to lipid rafts. Therefore, they have a fundamental function in for example neurotransmitter signal transduction. In contrast,

Flotillins/Reggies microdomains can also block signaling molecules, inhibiting interactions and stopping signaling responses (Allen et al. 2007).

Figure 1-4: Schematic representation of Flotillin-1 and Flotillin-2 in plasma membrane. Flotillin-1 and 2 are associated with lipid rafts in the inner leaflet of the plasma membrane through myristoylation (dark orange) and palmitoylation (green).

It has been proposed, through different studies on several cell types that Flotillin microdomains act as assembly locations for signaling that involve the activity of Src family kinases. For example, Flotillins in T cells are polarized to one side of the cell even in resting cells, and it has been shown that in addition to acting as signaling platforms, Flotillins also interact with specific phosphorylated proteins simultaneous with activation, which assist in identifying the immunological synapse location in activated T cells (Rajendran et al. 2003). Therefore, Flotillins may be involve in T cell receptor signaling because inducement of T cells with anti-CD3 and anti-CD28

antibodies leads to recruitment of the Linker for Activation of T cells (LAT) to Flotillin microdomains, which contain the Src family kinase Lck (Slaughter et al. 2003).

Flotillins/Reggies are also critical for signaling by GPI-anchored proteins and in the control of actin cytoskeleton reorganization, as well as actin-dependent cell adhesion and motility. A number of studies have confirmed the direct involvement of

Flotillins/Reggies in endocytosis. As the role of Flotillins/Reggies as signaling platforms, it is believed that they may accomplish their signaling role through endocytosis and interactions with the cytoskeleton (Babuke and Tikkanen 2007;

Langhorst et al. 2005b). Flotillin-1/Reggie-2 is involved in initiating cell-to-cell interactions (López-Casas and Del Mazo 2003), neuronal regeneration, insulin signaling in adipocytes (Baumann et al. 2000) and phagosome maturation in macrophages

(Dermine et al. 2001).

Von Philipsborn and co-workers (2005) studied the expression pattern of

Flotillin/Reggie proteins in different tissues in zebrafish during the early developmental stages. They suggested that Reggie proteins act as a functional platform for cell signaling in neurons and other cell types, leading to axon regeneration, cellular processes and contact formation. They also showed that Flotillin/Reggie proteins localize to the cell membrane, at cell-to-cell junction and along all early axon tracts.

They concluded that Flotillin/Reggie proteins are involved in neural development in zebrafish.

1.1.8 Lipid rafts and cell functions

It is believed that lipid rafts have a pivotal role in cellular functions and one of the principal functions of these microdomains is to compartmentalize cellular processes and 20

serve as organizing centers for signaling molecules and receptor trafficking (Korade and

Kenworthy 2009). Harder and colleagues (1998) hypothesized that lipid rafts can combine to form larger platforms, which concentrate signaling proteins so that these proteins can engage with each other in signal transduction processes.

The dispersed nature of these small-sized rafts is thought to be a central base for retaining these proteins in an inactive state as they are kept separate from each other

(Harder et al. 1998).

Lipid rafts have a vital role in many other different cellular processes including protein and lipid sorting and trafficking (Ikonen 2001), insulin-stimulated glucose transport (Chiang et al. 2001), cell adhesion and migration (Gómez-Moutón et al. 2001), bacterial and viral targeting of cells and toxins (Gruenberg 2003; Carrasco et al. 2004;

Smith and Helenius 2004), immune responses (Harder 2004), the stabilization of microtubules (Palazzo et al. 2004), axon growth and guidance (Kamiguchi 2006), cell apoptosis (Takahashi et al. 2006), and amyloid-β oligomerization (Hattori et al. 2006).

Furthermore, rafts are involved in many macrophage functions, such as endotoxin- mediated activation and cytokine production, major histocompatibility complex (MHC) class II antigen presentation (Korzeniowski et al. 2003) and cholesterol export (Gaus et al. 2004). Saltiel and Khan (2001) showed that lipid rafts have a role in substrate transportation, such as glucose and fatty acids, into the cell, implicating the localization of proteins, associated with substrate transportation, to lipid rafts (Saltiel and Kahn

2001).

1.1.9 Lipid rafts and disease

As lipid rafts are significantly involved in many cellular processes, such as membrane sorting and trafficking and cell polarization, they also have important roles in diseases since some pathogens use these microdomains to infect host cells (Van Der

Goot and Harder 2001). Influenza virus is one such virus that can use lipid rafts to infect cells (Scheiffele et al. 1999). This virus contains two integral glycoproteins, hemagglutinin and neuraminidase, that are considered raft-associated proteins. Influenza virus exits from the apical membrane of epithelial cells, which is rich in raft lipids, by budding and then spreads through the body (Zhang et al. 2000). Another example is

HIV-1, which integrates host raft lipids into its envelope, employing rafts to bypass the immune system and spread infection in the host cells (Alfsen et al. 2001).

A feature of Alzheimer’s disease (AD) is the development of plaques containing amyloid-β-peptide (Aβ), which is a derivative of amyloid precursor protein (APP), a large type I transmembrane protein (Selkoe 2001). Simons and Ehehalt (2002), hypothesized that APP is located at two different sites, one associated with raft regions at which Aβ is formed, and the other localized to non-raft regions, where α-cleavage occurs. They also showed the impact of cholesterol depletion on the distribution of APP from lipid rafts to the surrounding lipid bilayer. This study and others implied that there is a role of lipid rafts in AD.

The involvement of lipid rafts in disease pathogenesis has been investigated in a number of studies. One of these studies proposed that lipid rafts, particularly the caveolae subtype, may be involved in the pathogenesis of atherosclerosis, a blood vessel disease that is responsible for cardiac dysfunction (Shashkin et al. 2005).

Others looked for a correlation between raft microdomains and immune cell function, especially in T and B cells. Fan and his colleagues (2004) showed that there are changes in raft phospholipids of T and B cells under particular conditions (Fan et al.

2004). These alterations in lipid rafts constituents are thought to affect immune cell function by changing the number of receptors in lipid rafts (Li et al. 2005) as well as influencing the activity of some mediators such as AP-1 (activator protein 1).

Lipid rafts and some associated proteins are potentially involved in the pathogenesis of Parkinson's disease (PD), which is characterized by decreases in dopamine, in turn, which is involved in the control of movement, so that PP leads to uncontrolled muscle contraction and movement, as well as dementia (Tolosa et al. 2006), depression (Mentis and Delalot 2005) and anxiety (Richard 2005). One raft protein,

Flotillin-1, is up-regulated in the dopaminergic neurons of PD brains (Jacobowitz and

Kallaral 2004). Moreover, lipid rafts showed dynamic interaction with α-synuclein, a protein that has been implicated in PD pathogenesis (Eriksen et al. 2005). Other studies showed that α-synuclein co-localizes with CD55, a raft associated protein, and this protein depends on rafts to perform its function. As a result, it was concluded that there was a key role for lipid rafts in α-synuclein function and its role in PD (Fortin et al.

2004).

1.2 Methods to study membrane organization

1.2.1 Biochemical methods

Many studies have used detergent-based techniques for isolating lipid rafts such as

Triton X-100, a non-ionic surfactant detergent that dissolves cell membranes and

disrupts protein-membrane association, or other similar detergents. Detergent-resistant membrane fractions drift into the upper portions of sucrose gradients during ultracentrifugation and are differentiated as two or three opaque bands in the lighter density sucrose (Brown and Rose 1992).

Detergent-resistant membranes are an easy and useful method, and it was considered a powerful approach to determine the components of membrane rafts

(Schroeder et al. 1998). Despite this, the method suffers from some limitations. Firstly, the composition of the detergent-resistant membrane depends on the cell type and changing detergents and conditions yielded different results. As this method is performed at low temperatures, this will affect the organization of membrane order of the cell. Moreover, detergents such as Triton X-100 can alter the bilayer architecture

(Sot et al. 2002; Heerklotz et al. 2003). It is possible that detergents may extract lipids or proteins from within rafts themselves or extract non-raft lipids or proteins from the surroundings of rafts, which then creates different rafts, which do not resemble the original membrane domains (Shogomori and Brown 2003; Edidin 2003). Furthermore, since lipid rafts are linked to the actin cytoskeleton, detergents may disrupt cytoskeleton-raft interactions, creating separate subgroups of rafts (Hooper 1999; Janes et al. 1999). In addition, lipid rafts isolated with a detergent-based method may represent particular domains of the membrane, which are different from that prepared using non-detergent methods (Brown and London 1998; Pike et al. 2002; Eckert et al.

2003). In conclusion, detergent-based methods do not preserve the membrane structure.

Therefore, the isolated membranes do not represent actual raft domains as found in cell membranes. Nevertheless, this technique is still used in lipid rafts studies.

1.2.2 Advanced microscopy methods

Considering the drawbacks of the biochemical methods, it is thought that direct visualization would overcome these obstacles in studying and identifying lipid rafts, allowing better characterization and understanding of raft structure and function.

Many techniques have been implemented to study lipid rafts; most of these based on fluorescence microscopy because of the non-invasiveness of the techniques and because it can be applied to model and cell membranes. These techniques depend mainly on biophysical imaging methodologies such as fluorescence correlation spectroscopy (Bacia et al. 2004; Lenne et al. 2006), single molecule tracking microscopy (Kusumi et al. 2005), and fluorescence resonance energy transfer (FRET)

(Kenworthy et al. 2000; de Almeida et al. 2005). It was thought that these techniques would be useful for studying lipid rafts since they do not disrupt the cells. However, the need to label the desired proteins can affect and disrupt signaling cascades. Moreover, these approaches suffer from some drawbacks as the protein’s labels used may be too large, such that lipids do not pack properly into the membrane, thereby disrupting the rafts that they are investigating, or the labels may initiate lipid or protein crosslinking that can disturb membrane organization (Allen et al. 2007).

Fluorescence resonance energy transfer (FRET) microscopy, which is considered one of the most sensitive and selective tools for resolving spatial heterogeneity of molecular interactions within the cell (Sharma et al. 2004), is also utilized to study lipid rafts in intact cell membranes. In 1998, Kenworthy and Edidin implemented FRET with

<10 nm resolution to study raft microdomains enriched in GPI- anchored proteins in live cell membranes (MDCK). Their question was whether the GPI-anchored protein, 5′

nucleotidase (5′ NT), is randomly distributed in the apical plasma membrane of MDCK cells or whether it is assemble in raft microdomains. By detecting FRET under steady- state conditions, they observed that nearly all of 5′ NT molecules are randomly distributed and are not clustered due to lipid rafts. These outcomes were inconsistent with lipid rafts models and the membrane organization of GPI-anchored proteins

(Kenworthy and Edidin 1998). On the other hand, FRET was used to compare three different GPI-anchored proteins (folate receptor, CD59, and 5′ NT) and a glycosphingolipids (GSL) component of lipid rafts, in the plasma membrane of a number of different cell types. Cholera toxin B-subunit (CTxB) was used to label GSL

GM1. After detecting FRET between GSL GM1, labeled with (CTxB) and antibody- labeled GPI-anchored proteins, their data showed that those raft markers are in submicrometer structures within the plasma membrane. They concluded from their observations that lipid rafts either exist only in temporary stabilized structures, or when it is stable, it cover a minor portion of the cell membrane. These findings are consistent with the idea that a small portion of marker molecules are localized in small and transient rafts in the cell membranes, and that GSL and GPI-anchored proteins occupy only a small part of rafts. In contrast, it was revealed in previous FRET studies that most of GPI-anchored proteins and GSL are clustering in lipid rafts or with the presence of moderately large stable rafts (Kenworthy et al. 2000).

By combining FRET with electron spin resonance (ESR) data and spectral simulation, it has been found that (lo) and (ld) domains coexist in ternary mixtures of

1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC) and cholesterol together with either

1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) or 1-stearoyl-2-oleoyl-sn-

glycero-3-phosphocholine (SOPC). This study showed that these nanoscopic liquid domains in the ternary system behave nearly like the original phases, which may be useful in studying equilibrium thermodynamics (Heberle et al. 2010). FRET microscopy greatly assisted in illustrating the importance of phase diagrams of ternary lipid mixtures containing cholesterol, which has given great insight into cell membrane behavior. In a recent study, FRET provided new evidence that prokaryote membrane domains resemble eukaryotic lipid rafts, despite the differences in their structural difference in their sterols. By mixing different sterols with eukaryotic sphingolipids, the study showed that sterols can aid ordered domain formation, which is both necessary and sufficient for the formation of prokaryote membrane domains and sterols were also shown to be important for maintaining membrane integrity (LaRocca et al. 2013).

Fluorescence correlation spectroscopy (FCS), which is based on a correlation analysis of fluctuation of the fluorescence intensity, delivers a sensitive optical probe of the molecular dynamics in vivo and in vitro and it can detect a small number of molecules from nanomolar to picomolar concentrations (Chen et al. 2008). By using

FCS, Bacia et al. 2004 demonstrated the ability of FCS in recognizing a raft marker cholera toxin B subunit (CTxB) bound to gangliosides (GM1) and a nonraft marker

(dialkylcarbocyanine dye diI) in both live cell membranes and model membranes

(GUVs) through their different diffusional motilities. As the outcome of this study support the rafts existence in live cells, they proposed that fluorescence correlation spectroscopy is a promising tool to investigate the components of lipid rafts in live cells membranes (Bacia et al. 2004).

In previous studies, the binary mixtures of dimyristoylphosphatidylcholine/ distearoylphosphatidylcholine (DMPC/DSPC) were shown to form imperfect two-phase mixtures, which make it a suitable model for studying phase separation and lipid domain formation and for exploring the significance of FCS diffusion laws. The experimental FCS diffusion laws in both DMPC/DSPC mixtures and pure DMPC at distinct temperatures across the range of the phase transitions demonstrate apparent deviations from a pure Brownian motion. To clarify the cause of this deviation, numerical simulations were performed on these lipids. The outcome of this study indicated that both numerical and experimental methods showed remarkable similarities concerning the acquired FCS diffusion laws in terms of the effective diffusion coefficient and deviation from pure Brownian motion as a function of the lipid states.

Additionally, the study affirmed that FCS diffusion laws can be used to discriminate the existence of domains and define their mean size (Favard et al. 2011). Recently, FCS was implemented on a super-resolution STED microscope (STED-FCS) to measure the lateral diffusion of a new lipid analogue, which is a far-red emitting fluorescent phosphoglycerolipid analogue, to probe diffusion characteristics of lipids in membranes in the (lo) and (ld) phases. The authors detected micrometer size phases and found that, in both the (lo) and (ld) phases, the lipid analogue spreads easily on all spatial scales

(from 40 to 250 nm). This suggests that the rigid packing of the (lo) phase mainly reduces diffusion rather than causing uneven sub-diffusion. The data of this study advocate that STED-FCS in conjugation with the fluorescent lipid analogue may facilitate the study of lipid rafts in live-cell membranes (Honigmann et al. 2013). In the same year, FCS was applied with fluorescent phosphoglycerol and sphingolipid analogues to investigate the plasma membrane of living cells. At different 28

environmental conditions and on different lipid analogues, STED-FCS data disclosed molecular details of the observed nanoscale trapping. It also exposed big differences to the characteristics of (lo) and (ld) phase separation in model membranes (Mueller et al.

2013). Because of the narrow spatial resolution of conventional far-field optical microscopy, it is difficult to explore the molecular membrane dynamics in living cells, therefore, the superior spatial resolution of STED-FCS may help in making new findings in this field.

Single-particle tracking (SPT) is one of the techniques, which widely used in studying lipid rafts. This technique based on using colloidal-gold probes and optical trapping which allow moving the gold-particle-conjugated molecules or fluorophores in the living cell membrane. This technique offer a unique ability to observe the movement, gathering, and localization of individual single molecules in the plasma membrane of living cells in culture (Schmidt and Schütz 1995; Saxton and Jacobson

1997; Sako et al. 2000).

Pralle with his collaborators estimated the local diffusion of single membrane proteins in the membrane of living cells by high resolution single particle tracking. They observed that once GPI-anchored and transmembrane proteins are localized in raft region their diffusion is significantly decreased compared with that of nonraft transmembrane proteins. Also, they revealed that cholesterol depletion increases the diffusion of raft-linked proteins for transmembrane raft proteins to the level of transmembrane nonraft proteins and even more for GPI-anchored proteins. They estimated that the range of rafts sizes is from 50 to 200 nm in diameter (Pralle et al.

2000).

In a related study, Dietrch et al. in 2002 investigated in depth the physical and chemical properties of transient confinement zones (TCZs) and their relevance to rafts using SPT. They discovered that cholesterol depletion significantly decreased zone redundancy, and the diffusion within the zones was reduced by a factor of ∼2. As a result, they proposed that a main cause of the observed transitory zone size is due to the existence of raft domains. The data presented in this study indicated that there are raft associated domains localized in definite regions of the plasma membrane of murine fibroblasts cells, which have a lifetime of tens of seconds (Dietrich et al. 2002).

The drawback about SPT is that gold or latex particles are used as probes; therefore, there is a chance of probe-induced crosslinking. Even so, Kusumi and his colleagues strongly recommend using single-molecule techniques, particularly high speed SPT, because of the valuable contribution that technique offers in investigating raft dynamics, structure, size, and lifetime (Kusumi et al. 2005).

Atomic force microscopy (AFM), which is not a fluorescence technique, is considered a powerful approach to identify membrane order at high resolution and is able to detect the greater thickness of the bilayers in model liquid-ordered versus - disordered membranes. This allows investigators to perform experiments on biological specimens and can produce images under near-physiological conditions (Henderson et al. 2004). AFM also provided informative data from living cells which helps in understanding the molecular mechanisms of cell function (Roduit et al. 2008). AFM still suffers from limited spatial resolution in live cell imaging (Shi et al. 2012) but despite its superior resolution, it has some disadvantages when used to study supported bilayers, which may show non-physiological behaviors under particular conditions

(Rinia et al. 2001; Yuan et al. 2002). In addition, different membrane structures cannot to be resolved in situ using AFM (Henderson et al. 2004). For example, when AFM was applied to erythrocyte membranes, although it detected the structure of the erythrocyte membrane skeleton, it could not analyze the rafts because of the rough erythrocyte surface (Takeuchi et al. 1998).

1.2.3 Environmentally sensitive dyes

Since fluorescence microscopy is a reliable option to investigate membrane lipid domains, a probe that alter its fluorescence properties based on the polarity of its surrounding environment and has the ability to partition into the ordered and disordered domains, is required for best results (Parasassi et al. 1998). Therefore, a straightforward way to study lipid rafts is to use environmentally sensitive membrane dyes (Gaus et al.

2003; Jin et al. 2006; Owen et al. 2012).

Laurdan (6-dodecanoyl-2-dimethylaminonaphthalene) is considered one of most convenient polarity-sensitive dyes, which was produced from Prodan by Weber and

Farris in 1979 (Weber and Farris 1979). It has been used successfully to study bilayer properties in both model and cellular membranes because of its sensitivity to membrane phase transitions (Henshaw et al. 1998; Harris et al. 2001). Laurdan integrates into the cell membranes and aligns itself in parallel with the phospholipids’ hydrophobic tails (Bagatolli et al. 1998), and it displays spectral sensitivity to the polarity of its environment with a ∼50-nm red shift of its emission maximum wavelength in polar versus nonpolar environments. Hence, fluorescence intensity measurements at two properly selected wavelength regions provide information on the membrane polarity.

The study showed that Laurdan spectroscopic properties reports topical water content in the membrane (Parasassi et al. 1995) (Figure 1-5).

Figure 1-5: Schematic of the emission shift in polar and non-polar environments. A) In a solution environment, solvents surrounding the ground state fluorophore have dipole moments that can interact with the dipole moment of the fluorophore to produce an ordered distribution of solvent molecules around the fluorophore. The differences in energy level between the ground and excited states in the fluorophore yield a change in the molecular dipole moment, which eventually prompts a reordering of surrounding solvent molecules. After the fluorophore has been excited to higher vibrational levels of the first excited singlet state (S(1)), extra vibrational energy is quickly lost to surrounding solvent molecules as the fluorophore slowly relaxes to the lowest vibrational energy level. Solvent molecules facilitate stabilizing and further decreasing the energy level of the excited state by re-orienting (solvent relaxation) around the excited fluorophore. This has the effect of lowering the energy separation between the ground and excited states, which results in a red shift of the fluorescence emission. The polarity of the fluorophore also defines the sensitivity of the excited state to solvent effects. Polar and charged fluorophores show a stronger effect than non-polar fluorophores. B) Laurdan is excited at 800 nm (red arrow). The dye fluoresces with a peak emission wavelength around 450 nm (violet) when exist in the ordered phase and ~500 nm when exist in the disordered phase (blue) with an emission shift around 50 nm.

Generalized polarization (GP) measures the relative water content of the membrane and the data is calculated with the following formula: GP = (I440 - I490)/ (I440 + I490), where I440 and I490 are the emission intensities at the blue (440 nm) and red (490 nm) channels of the emission spectrum that represent the fluorescence emission maxima in

the ordered and disordered phases, respectively. In model membranes, it was observed that the GP distribution in the ordered phase vesicles is quite narrow while the GP distribution in the disorder phase vesicles is broad (Parasassi et al. 1997). The Laurdan dye has some limitations such as rapid photobleaching, although the development of the two-photon fluorescence microscopy has made it possible to minimize this (Parasassi et al. 1997).

In general, Laurdan can be applied to model membranes, live and fixed cells, and it has been found to be a suitable environmental dye to study lipid order in cells, even when it is used in whole organisms, Laurdan did not show any adverse effects on the membranes (Owen et al. 2010; Dinic et al. 2011; Owen et al. 2012).

In 1997, two-photon microscopy with Laurdan demonstrated that there were regions of differential membrane fluidity in phospholipid vesicles, red blood cells, a renal tubular cell line, and in purified apical renal brush borders and basolateral membranes. Vesicles of an equimolar mixture of 1, 2-Dioleoyl-sn-glycero-3- phosphocholine (DOPC) and 1, 2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) showed coexisting rigid (ordered) and fluid (disordered) domains; however, the rigid domains still have GP values that are lower than pure gel-phase domains, suggesting that the rigid domains are still fluid membranes. By mixing DOPC/DPPC with 30% mol cholesterol, their GP distribution became narrower, due to the cholesterol incorporation

(Parasassi et al. 1997). These experiments confirmed that the domain morphology depends on the presence of cholesterol.

In another study the fluorescent probes Laurdan, prodan, and N-Rh-DPPE were used to detect domain coexistence in GUVs. This study showed that Laurdan was

preferred over other probes such as prodan and N-Rh-DPPE because it partitioned more equally and had higher photoselection than other probes (Bagatolli and Gratton 2000).

Thus, the heterogeneity and phase separation obtained using Laurdan in combination with two-photon microscopy provide an effective tool to study the properties of the membrane system.

The efficiency of using Laurdan GP was verified by Harris and co-workers (2002),

showing that GP and polarization techniques could improve the understanding of

membrane phase properties. The phase transition from the ordered phase to the

disordered phase was characterized by a decrease in the Laurdan GP value. Based on

these results, they proposed that Laurdan anisotropy could reveal alterations in

membrane fluidity while Laurdan GP detected alterations in phospholipid order.

Phospholipid order means the order parameter, liquid-ordered and liquid-disordered

phases, and it is related to the configurations of the lipid tails or to the conformational

order of phospholipids based on the acyl chains. Fluidity is related to the diffusability

of lipids in the bilayer, it reveals the orientational order of whole or partial acyl chain

of the phospholipids and the rotational and lateral diffusional motion of the acyl chain

(Xingjie and Youguo, 2001). Fluidity is affected by the presence of cholesterol, the

phospholipid composition, and the length and degree of unsaturation of the

phospholipid fatty acyl chains (Owen et al. 1982). Therefore, one could not be

changed without affecting the other; for example, if cholesterol is added to the

membrane it will reduce the fluidity and increase the membrane order simultaneously,

thus any changes occurring to fluidity will affect the phospholipids order and vice

versa. Consequently, the advantages of using Laurdan in the model and cell

membranes was improved by inclusion of a polarized excitation source (Harris et al.

2002). The disadvantage of using a polarized excitation source (laser) is that it

preferentially excites molecules when the dipole is in the same direction as the

polarization, which means that it can give a wrong answer between vertical and

horizontal membranes especially if the dyes are held rigidly. For example, in the gel

phase, there is a strong photoselection effect as rotational freedom of the fluorophores

is limited while a dye is in the liquid phase, where lipid order is low, is excited

strongly since the fluorophore can rotated and be aligned parallel to the excitation

polarization (Parasassi et al. 1997). As a result, there is a difference in the emission

intensity between the parallel and perpendicular orientations of the dye transition

dipole.

Direct visualization of membrane lipid structures in living cells can also be carried out using two-photon microscopy with Laurdan. By comparing the properties of microscopically visible domains with those of isolated detergent-resistant membranes, membrane regions of high order were indirectly linked to lipid rafts. But it has been proposed that GP values of living cells and model membrane may not match with each other. Moreover, cholesterol was shown to play a vital role in existence of highly ordered cellular membranes (Gaus et al. 2003). In this study, Laurdan, through constructing GP images, facilitate investigating lipid rafts in living cells which may provide another evidence that support lipid rafts hypothesis.

The di-4-ANEPPDHQ probe can be used with fluorescence-lifetime imaging microscopy (FLIM) and applied to model membranes and live cells (Owen et al. 2006).

The authors confirmed the ability of the probe to visualize liquid-ordered phases in vivo

by extending the technique to live cell imaging of epithelial cells. The dye was able to detect varying levels of ordered phase in live epithelial cells. Jin and colleagues (2006) employed di-4-ANEPPDHQ dye together with both single photon fluorescence and two photon fluorescence to image the phase separation in giant unilamellar vesicles (GUVs).

These studies showed that FLIM and GP measurements were verified to report high and low membrane order in model membranes (Jin et al. 2006).

Kim and his colleagues developed a variant of Laurdan, 6-dodecanoyl-2-[N- methyl-N-(carboxymethyl) amino-naphthalene or C-Laurdan. They thought that C-

Laurdan could outperform Laurdan as it had a greater sensitivity to membrane polarity, a brighter two-photon fluorescence signal and remained in the plasma membrane for longer than Laurdan. C-Laurdan was used with two-photon microscopy to visualize and image lipid rafts. The study showed that lipid rafts cover approximately one third of the cell surface and that C-Laurdan was useful as a reliable two-photon probe for cell membrane imaging (Kim et al. 2007). However, this probe is not commercially available and is difficult to prepare.

Further optimization was performed on C-Laurdan by designing 6-[(E)-3-oxo-1- dodecenyl]-2-[N-methyl-N-(carboxymethyl) amino-naphthalene (CL2). CL2 has a longer conjugation length, improving the two photon cross-section for a brighter two- photon microscopy imaging and enhancing the sensitivity of the fluorescence intensity relative to its environment polarity for the selective detection of the lipid rafts in live cells and tissues without relying on GP values. By using two-photon microscopy, it was shown that this probe may provide direct visualization of lipid rafts in the live cells and the pyramidal neuron layer of the CA1 region at a depth of 100-250 µm in live tissues

(Kim et al. 2008). However, the influence of this kind of probe on intact organisms and what effects it may have on the organism viability is unknown.

Owen and co-workers (2010) reported the first image of membrane lipid order in a whole, living vertebrate organism. Images of cell membranes in tissues of live zebrafish embryos were taken in three dimensions using Laurdan and multiphoton microscopy. A high lipid order in the apical surfaces of polarized epithelial cells was observed in this study (Owen et al. 2010).

1.3 Lipid rafts debates

Many studies were performed exploring lipid rafts to clarify their size, structure, and their involvement in cell functions. It is difficult to study lipid rafts in living cells because of the resolution limitation of conventional microscopy techniques. Such limitation has occurred because of the wave transmission nature of light. For instance, diffraction makes a tiny object look bigger than its actual size. Lipid rafts have been estimated to be around 20 nm in size, which is under the resolution limit of ~ 200 nm, therefore, more advanced techniques are needed for more accuracy in raft imaging.

There is also no consensus between different studies regarding the size of lipid rafts in the cell membrane. However, it should also be noted that it is currently not know how many molecules are typically within a raft for any given time period.

Different methods are used to study lipid rafts, including detergent extraction and using cholesterol depletion to disrupt lipid rafts in living cells. Most of these studies have been carried out on model membranes, which may not reflect real cellular membrane. While biophysical studies on model membranes show that the existence of

liquid-ordered and liquid-disordered microdomains is physically possible, there are some concerns in applying this model to living cells.

Cellular membranes contain a heterogeneous mixture of lipids with many proteins that can interact with membrane lipids. Phase segregation is influenced by temperature, the lipid and protein composition and lateral pressure. Depending on these conditions, model membrane studies may not demonstrate what truly happens in cellular membranes because these studies are typically performed with one or two homogeneous phospholipids at non-physiological temperatures (Munro 2003).

Cholesterol depletion is also a common method to study lipid rafts. In this method, cells or model membranes are incubated with cyclodextrin, which extracts cholesterol from the membranes. This method also has drawbacks as it can also reduce the lipid, PI

(4,5)P2, which has an essential regulatory function in the cell (Pike and Miller 1998).

Moreover, cyclodextrin may extract cholesterol from raft and non-raft domains, and it may remove phospholipids from the cell membrane along with modifying the distribution of cholesterol between plasma and intracellular membranes (Zidovetzki and

Levitan 2007). Others have indicated that disruption of PI (4,5)P2 (PIP2) has a similar outcome to cholesterol depletion. Therefore, a conclusion came that the loss of any cell function as a result of cholesterol depletion may not be due to raft disruption as non-raft processes may also be affected (Kwik et al. 2003).

The role of PIP2 in cell morphology, motility, endocytosis, exocytosis, phagocytosis, and T cell activation, in addition to the association of PIP2 with membrane rafts, have been investigated. Different models of PIP2 signaling from the plasma membrane to the actin cytoskeleton were performed to understand the regulation mechanisms in different

cellular functions. It has been proposed that PIP2 mediates signaling by concentrating

PIP2 and hence PIP2-binding proteins into membrane rafts via binding of raft-associated proteins (Johnson and Rodgers 2008). In an unusual case, Milhiet and co-workers

(2002) observed that lipid microdomains in the renal and intestinal brush border membrane can form in the absence of cholesterol and may disappear in the presence of high levels of cholesterol. They also propose that cholesterol depletion does not disrupt lateral heterogeneity in intestinal and renal brush-border membranes (Milhiet et al.

2002).

Douglass and Vale (2005) used single-molecule imaging techniques to study the dynamic properties of the signaling network and found that these microdomains are formed by protein-protein interactions and not through lipid rafts. On the other hand, there was no indication that the signaling abilities of these molecules are retained in the absence of lipid rafts (Douglass and Vale 2005). However, in vivo experiments are required to explain and verify these findings. Microscopy techniques such as fluorescence microscopy may reveal more details about lipid rafts regardless of the diverse results. Despite the different techniques showing the involvement of lipid rafts in cellular processes, there is still more work to be done to resolve these issues (Munro

2003). In addition, until now little is known about rafts in cells when cells are still part of tissue, which also need to be investigated.

1.4 Organization of polarized epithelial cells

1.4.1 Physiological function of polarized epithelial cells

Polarized epithelial cells perform several significant biological functions in different tissues such as protection from outer environment, secretion, and absorption of nutrients, and transportation of biological materials. They act as a barrier to cover the inner and outer linings of various body cavities, which protect the underlying tissues from physical trauma, infection, and toxins. In addition, polarized epithelial cells regulate material exchange between the underlying tissue and the body cavity, such as in the intestine by using active transport systems, which are located on the apical region of the plasma membrane. Certain epithelial cells have the ability to use active transport systems to absorb filtered materials, which can then be distributed to the rest of the body.

All cells are capable to endocytosis essential materials for cell growth and signaling.

Epithelial cells are responsible for the secretion of fluids and compounds that are necessary for processes such as secretion of hormones, which are involved in development and reproduction, into the circulation. In addition to secretion of waste products such as sweat, mucus to lubricate body cavities, enzymes for digestion and for protection, and other products that regulate different metabolic processes in the body

(Alfons and Lommel 2003). A key feature of polarized epithelial cells is the ability to arrange their plasma membrane into structurally and functionally discrete compartments, which is necessary for the ordinary physiological function of epithelial cells.

The polarity feature of epithelial cells is responsible for various biological processes. One of these functions is to establish selective permeability barriers of

various ionic compositions between the lumen and serosa. Another important task is the specialized vesicular transport, which relies on polarized deliveries of specific enzymes to the apical and basolateral membranes, e.g., during absorption, transcytosis, and secretion, allowing regulation of the composition of the lumen and serosa (Berridge

1972; Simons and Fuller 1985).

1.4.2 Membrane organization in polarized epithelial cells

Many in vivo and in vitro studies have confirmed the fundamental features of polarized epithelial cells. These cells are subdivided into three discrete surface regions - the apical, lateral, and basal – that are all part of the plasma membrane. The apical and basolateral membranes are separated by tight junctions, which regulate the diffusion of internal macromolecules and ions and prevent the mixing of proteins and lipids between the apical and basal membranes. The apical and basolateral membranes have different lipids and proteins compositions as well as different functions (Simons and Meers 1988;

Simons and Ikonen 1997). The apical membrane is robust because it is mostly enriched with high levels of sphingolipids and cholesterol which form a packed membrane environment, while the basolateral membrane is enriched in glycerolipids and phosphatidylcholine (Rodriguez-boulan et al. 1989). It has been shown that the apical and basolateral plasma membranes of MDCK cells have distinct phospholipid compositions. This was recognized when MDCK cells were labeled with [32P] orthophosphate until the phospholipid groups were equally labeled then the labelled

MDCK cells were infected with fowl plague virus (FPV), an avian influenza virus, or with vesicular stomatitis virus (VSV). The viruses budding were isolated from either domain, and its phospholipid compositions were analyzed. The results showed that their

phospholipids are significantly different. Therefore, it was assumed that the two viruses acquired their lipids from the plasma membrane in a similar way and that the different phospholipid compositions of the viruses from polarized cells exposed differences in the phospholipid composition of the apical and basolateral membranes (Meer and Simons

1982).

The apical membrane faces the lumen and its exterior leaflet is enriched in glycosphingolipids, while the interior leaflet is enriched in phosphatidylinositol-4, 5- bisphosphate (PtdIns (4,5)P2) (Martin-Belmonte et al. 2007). Lateral membrane consists of cell adhesion molecules, junctional complexes, such as desmosome and zonula occludens, and tight junctions mediate cell-to-cell contacts and interactions.

Junctional complexes also contains the lipid phosphatidylinositol-3,4,5-

triphosphate (PtdIns (3,4,5)P3) (Gassama-Diagne et al. 2006). The basal membrane contains basement membrane receptors and is in contact with the extracellular matrix

(ECM) (Rodriguez-boulan et al. 1989; Alberts et al. 2002).

The asymmetric distribution of proteins and lipids in the apical and basolateral membranes is responsible for the apical–basolateral polarity in epithelial cells, which is a hallmark of epithelial cells. The membrane and secretory proteins are distributed to certain regions on the plasma membrane, creating membrane asymmetry, which maintain cell functions such as membrane trafficking (Mostov et al. 2003).

1.5 How is the membrane organization in polarized epithelial cells maintained?

1.5.1 Lipid rafts in polarized epithelial cells

It has been hypothesized that lipid rafts in epithelial cells localize to the apical domain of the plasma membrane so that apical membranes are more ordered than basolateral membranes (Simons and Meers 1988). Moreover, it has been proposed that lipid rafts also exist in the biosynthetic and endocytic pathways. Cholesterol is synthesized in the endoplasmic reticulum (ER) while sphingolipids are synthesized and their head-group modified in the Golgi (Meer 1989). It was observed that lipid rafts first assemble in the Golgi (Brown and London 1998) and are then delivered towards the plasma membrane. Vesicles returning back to the endosomes from the plasma membrane contain little sphingomyelin or cholesterol (Brügger et al. 2000). In several cell types, the incorporation of proteins into rafts is essential for polarized delivery to the cell surface (Simons and Ikonen 1997; Keller and Simons 1997).

The sorting of many apical proteins may be directed by lipid-lipid and lipid-protein interactions. The enrichment of sphingolipids in the apical membrane, together with their propensity to associate with cholesterol to form lipid rafts, has directed to the concept that rafts favorably traffic to the apical membrane after intracellular assembly.

Since different proteins incorporate with rafts during apical transport process, rafts might act as apical sorting platforms (Simons and Ikonen, 1997; Klemm et al. 2009;

Surma et al. 2012).

The establishment and maitenance of cell polarity is critical for tissue development and normal physiological cell function. An inability to establish or maintain cell polarity 43

alters the function of epthelial cells which leads to disease such as hypercholesterolaemia and kidney disease (Mellman and Nelson 2008).

To establish and maintain the polarized phenotype of epithelial cells, trafficking and sorting mechanisms that deliver proteins and lipids to their correct domains in the plasma membrane are required. At the cellular level, the establishment and maintenance of cell polarity have similar mechanisms in different tissues and organisms, and the main characteristic of cell polarity is the asymmetric organization of the plasma membrane. This asymmetry is mostly accomplished through membrane trafficking that is under the control of signaling molecules such as the Rho family of small GTPases

(McCaffrey and Macara 2009; Nelson 2009).

1.5.2 Membrane trafficking

It has been proposed that membrane trafficking has major contributions to the maintenance of cell membrane asymmetry (Orlando and Guo 2009). In membrane trafficking, the newly synthesized membrane proteins, and lipids are transferred from the endoplasmic reticulum (ER) to the Golgi and then co-transported to the trans-Golgi network (TGN) where these proteins are sorted and delivered to their particular destinations in the plasma membrane (Griffiths and Simons 1986; Thiele and Huttner

1998; Mellman and Nelson 2008). In polarized epithelial cells, three different trafficking pathways are involved in delivering proteins from the donor membrane to the apical and basolateral membranes The first pathway acts via direct protein delivery from the TGN to the apical or basolateral membrane through exocytosis, secretion, or anterograde trafficking. In the second pathway, proteins are delivered to the plasma membrane to be processed or to control their activity through endocytosis.

The third pathway involved relocating proteins from their position on the plasma membrane to a different location via transcytosis (Mostov et al. 2000; Peer 2011).

During membrane protein transportation from the ER to the Golgi and to the plasma membrane, membrane proteins undergo an important modification in these compartments, namely, the processing of complex oligosaccharide chains. Vesicular structures act as intermediates, which transport these membrane proteins to their destination. Three classes of protein complexes that form these vesicular intermediates have been identified (Lee et al. 2004; Pearse and Robinson 1990; Rothman 1994): coatomer protein complex-II (COPII) complexes are essential for ER-Golgi trafficking,

COPI complexes which are essential for Golgi-ER trafficking and intra-Golgi transport, and the adaptor protein (AP)-clathrin complex which is important for transport between the Golgi, the plasma membrane, and endosomes. Protein sorting involves distinguishing sorting signals and recruiting different cargo proteins into transport vesicles through the particular proteins of each complex, while the unrecognized membrane proteins are omitted from the vesicles. However, other complex proteins distort and sculpt the membrane to produce the membrane vesicle. The different AP complexes are localized to different membrane compartments to grab discrete sets of cargo proteins before directing them to their target. The AP-clathrin complex also has additional proteins that increase the complexity of cargo-selection approaches and affect membrane curvature (Edeling et al. 2006). Actin and the microtubule cytoskeleton facilitate vesicle trafficking between compartments (Hehnly and Stamnes 2007) and the organelle-specific Rab GTPases, while the vesicle-tethering complexes regulate membrane trafficking by targeting secretory vesicles to specific domains of the plasma

membrane (Grosshans et al. 2006) and Soluble NSF Attachment Protein Receptor

(SNARE) assembly controls vesicle fusion with the target plasma membrane (Rothman

1994).

The above mechanisms are considered the key of membrane trafficking in all cells but in polarized cells they are adjusted to sort proteins into distinct plasma membrane domains. Once these proteins are localized to their correct destination they are separated from each other by tight junctions so that the apical polarity proteins cannot enter the basolateral domain and vice versa (St Johnston and Ahringer 2010).

1.5.3 Polarity proteins

While different types of proteins are found at the apical and basolateral membranes, only three key protein complexes are thought to be required to establish and maintain cell polarity in epithelial cells. These highly conserved polarity proteins complexes are: the Crumbs complex contains crumbs (Crb), PATJ (protein associated with tight junctions), and Pals1-associated tight junction protein, which is localized to the apical membrane adjacent to the junctional complex. The second polarity complex is the partitioning defective (Par) complex contains atypical protein kinase C (aPKC), Par3,

Par6, scaffold proteins, and the Rho family GTPase Cdc42, which is also localized to the apical membrane adjacent to the junctional complex like the Crumbs complex

(Figure 1-6). The third complex is Scribble, which contains Scribble (Scrib), Discs

Large (Dlg), and lethal giant larvae (Lgl), and is localized mainly under the tight junctions within the basolateral membrane and promotes basolateral polarity (Bryant and Mostov 2008).

Atypical protein kinase C (aPKC) has two isotypes, aPKCζ and aPKCλ, both of which are involved in signaling through lipid metabolites such as phosphatidylinositol

3-phosphates. ASIP (aPKC-specific interacting protein), the first vertebrate homologue of a Par protein, was recognized as the orthologous of C.elegans Par3. Furthermore,

ASIP and Par3 share three PDZ domains, and both have the ability to bind to aPKC

(Izumi et al. 1998). The PAR complex can also be divided into two subgroups: CDC42–

Par6–aPKC which associated with the apical membrane and Par3–aPKC, which also associated with tight junctions (TJ) and is involved in delivering the phosphatase and tensin homolog (PTEN) to the tight junctions. Both Crumbs and Par complexes establish cell polarity through their significant involvement in the assembly and function of the junctional complex, which acts as a regulator of molecule transportation between the apical and basolateral membranes.

In summary, the three complexes, Crumbs, Par, and Scribble, regulate the formation of tight junction and epithelial polarity through interacting with each other. The Crumbs complex is responsible for organizing the formation of the apical surface, while the

Scribble complex controls the lateral membrane establishment, and the Par complex equalizes between apical and lateral membranes through multiple interactions

(Pieczynski and Margolis 2011)

Figure 1-6: Schematic representation of localization of Par & Crumbs complexes. Polarized epithelial cells have two distinct membranes, apical membrane (red) and basolateral membrane (green), and each of them has a different composition and different functions. Polarity proteins Par and Crumbs are localized adjacent to tight junction at the apical membrane of polarized epithelial cells.

In zebrafish, the crb3a and crb3b genes, which are orthologous of mammalian

CRB3, encodes two different proteins, Crb3a and Crb3b, of 109 and 96 amino acids, respectively. Crb3a is highly expressed in the otic vesicle, and both Crb3a and Crb3b are expressed in the digestive tract primordium. Crumbs genes in zebrafish show various expression patterns. For example, crb1 and crb2b genes are mostly expressed in the central nervous system, in the brain, in the photoreceptor cells in the retina, and in the pineal gland (Omori and Malicki 2006). Mutation data led to the proposal that crumbs genes have a pivotal role in zebrafish development. In general, crumbs genes in invertebrate and vertebrate models also show various expression patterns (Klebes and

Knust 2000; Izaddoost et al. 2002).

Pard3 is the Par3 ortholog in zebrafish and it encodes two Pard3 proteins (150 and

180 kDa) that differ in their carboxyl-terminus. It has been proposed that Pard3 is localized to the apical region of the retinal and brain neuroepithelium cells in zebrafish

(Wei et al. 2004).

The aPKC complex, which is composed of aPKC, Par3, and Par6, plays a central role in the establishment and maintenance of epithelial cell polarity. These three proteins function as one element by interacting dynamically with each other to regulate their asymmetric localization at the cell membrane (Suzuki and Ohno 2006; Ohno 2001;

Macara 2004). The aPKC complex is localized asymmetrically in different cells. In C. elegans, at the one-cell embryo stage, the aPKC complex is localized to the anterior cortex (Hung and Kemphues 1999; Tabuse et al. 1998) and in Drosophila, it is located at the apical surface of embryonic epithelial cells and the neuroblast (Petronczki and

Knoblich 2001). In mammals, the aPKC complex is localized asymmetrically together with the ZO-1 protein at the TJ (tight junction) of epithelial cells (Izumi et al. 1998;

Suzuki et al. 2001). Researchers have found that defects in any one of those three components, aPKC, Par3, or Par6, may lead to disturbances in the asymmetric localization of all other Par proteins, suggesting that the localization of these proteins is mutually dependent (Ebnet et al. 2001; Itoh et al. 2001). As a result, it has been suggested that aPKC complex activity is needed to originate its asymmetric distribution.

Interestingly, it has been shown that the function of the aPCK complex is disrupted when the aPKC phosphorylation site in the Par3 protein is mutated (Tabuse et al. 1998).

Furthermore, researchers have noted that Par3 shows a slightly distinct localization from aPKC and/or Par6 in polarized epithelial cells (Harris and Peifer 2005). Others

showed that aPKC and its kinase activity are essential for the establishment, rather than maintenance, of cell polarity in epithelial cells. Suzuki et al. (2001) overexpressed a dominant-negative mutant of aPKC (aPKCkn) in MDCK II cells to observe the effect of this defect on cell polarity. This mutation led to mislocalization of ASIP/Par3 and Par6 proteins. It also caused a mis-localization of the tight junction proteins ZO-1, occludin, and claudin-1 as well as disrupted the barrier function of the tight junctions without affecting cell-to-cell contact.

As a result, mis-localization of apical and basolateral membrane proteins occurred.

Interestingly, it has been observed that the aPKC mutation only affected cells that are developing cell polarity without any effect on fully polarized cells. Therefore, it was proposed that aPKC kinase activity is important for the establishment, rather than the maintenance, of cell polarity. However, the kinase activity of aPKC and the binding of

Par6 to Cdc42 have been described to play an essential role in maintaining Par3 localization at the cortex (Hutterer et al. 2004). It has been assumed that Par3 directs the aPKC–Par6 complex to particular membrane regions since it was observed that Par3 frequently reaches its destination earlier than aPKC and/or Par6 and can remain there even if aPKC or Par6 are depleted (Harris and Peifer 2005; Tabuse et al. 1998).

The Crumbs (Crb) complex, a conserved protein complex localized to the apical membrane, is a key regulator that maintains epithelial cell polarity through its pivotal cooperation with the aPKC (Hurd et al. 2003). It has been shown that Crb plays an influential role in stabilizing the subapical localization of aPKC during cellular shape changes such as late gastrulation. On the other hand, the aPKC complex also is necessary for the stable apical localization of the Crb complex and aPKC is even more

critical than Crb in epithelial cell polarity (Bilder and Perrimon 2003; Tanentzapf and

Tepass 2003). Gibson and Perrimon (2003) confirmed that the Crb complex stabilizes the apical localization of aPKC without interacting with Par3 due to insufficient evidence of the interaction of Crb complex with the LgI group (Gibson and Perrimon

2003).

The Crumbs complex might also be responsible for the development of apical membrane identity after the completion of junctional maturation because in Drosophila,

Crumbs mediates the recruitment of the actin-binding protein D moesin and the components of the apical spectrin-based membrane skeleton (Medina et al. 2002). It has been found that mutations in one of Crumbs complex components are associated with retinal degeneration in humans, mice, and flies. The Crumbs complex is also proposed to play a role in the development of other disease processes that are based on epithelial dysfunction, such as tumorigenesis (Bulgakova and Knust 2009).

1.6 Zebrafish as a model to study epithelial cells in vivo

1.6.1 Introduction to zebrafish

Zebrafish has a great potential in life science and has become an ideal model organism for the study of vertebrate development and disease, functional genomics, organ function, behavior, toxicology, and drug discovery. It is considered one of the most important vertebrate model organisms in the life sciences (Vascotto et al. 1997;

Grunwald and Eisen 2002; Amsterdam and Hopkins 2006).

Zebrafish, which is a popular aquarium fish, is a tropical freshwater fish that belongs to the cyprinid family, along with minnows and carps. Zebrafish are classified

as omnivorous fish, and in the wild, it mostly feeds on zoo- and phyto-plankton and insects. In the aquarium, zebrafish are usually fed different types of dry food (with a fat content of 8–15%) along with live food, most commonly Artemia (brine shrimp) that are rich in cholesterol and unsaturated fatty acids. Zebrafish start to feed at around 5 dpf

(days post-fertilization); until that time, the only source of energy is the maternally derived yolk (Brand et al. 2002).

Zebrafish have many advantages over other experimental animals. It is durable fish, considerable numbers of fish can readily be raised at low-cost, and they breed all year round. Females can spawn every 2–3 days and a single clutch may contain several hundred eggs. Relative to other fish egg sizes, zebrafish eggs are larger (0.7 mm in diameter at fertilization), which make microinjection procedures easier to perform

(Kimmel et al. 1995). Embryos develop outside the mother and they are transparent which make them ideal for light microscopy. Zebrafish can grow up to 6.4 cm and their lifespan in captivity is around 2–3 years but may extend to 5 years under ideal conditions (Spence et al. 2008). Zebrafish have a short development period; embryos develop from the egg to larvae stage in less than 3 days, with precursors to all main organs evolving within 36 h. Within 5 dpf, larvae display food seeking and active avoidance behaviors (Kimmel et al. 1995). Moreover, zebrafish development is well understood and can be traced easily, beside its steady size during early development facilitates straightforward techniques such as staining as the dye may administered by adding it directly to the fish container (Spence et al. 2008).

The transparency of zebrafish embryos together with new imaging techniques enables non-invasive visualization of organs with high resolution imaging of the whole

organism, which is preferred by many researchers around the world. Many fluorescent lipid analogues and probes, in conjunction with various microscopy techniques, have been successfully implemented in zebrafish studies (Hölttä-Vuori et al. 2008; Stoletov et al. 2009). New nonlinear optical techniques include label-free visualization of lipids and open new research opportunities using zebrafish as a research model in lipid studies

(Hölttä-Vuori et al. 2010).

1.6.2 Structure and function of polarized epithelial cells in zebrafish

Polarized epithelial cells are the basic building blocks of different ductal organs, such as the kidney, gut, and liver, in most living organisms. These cells have attracted the interest of researchers due to their vital role in health and disease.

It has been observed that the zebrafish gastrointestinal system, which includes the liver, pancreas, gall bladder, is a linearly segmented intestinal track with absorptive and secretory functions, closely resembling that of mammals. Zebrafish has no stomach and the intestinal epithelium of zebrafish exhibits proximal-distal functional specification and has similar epithelial cell lineages found in mammals such as absorptive enterocytes, goblet cells, and enteroendocrine cells (Ng et al. 2005; Wallace et al. 2005). For example, enterocytes in zebrafish have basolateral nuclei and form tight junctions, and have apical microvilli and an intestinal brush border like those in mammals (Detrich et al. 2010).

The intestine of the zebrafish consists of a long intestinal epithelium that folds twice in the abdominal cavity. This intestinal epithelium is separated into three different sections: the intestinal bulb, the mid-intestine, and the posterior intestine. The abundant intestinal folds gradually become shorter in a rostral-to-caudal direction as the posterior 53

intestine is an unfolded monolayer of simple columnar epithelium. These folds are significantly larger than the finger-like intestinal villi of mammals and the columnar- shaped absorptive enterocytes and goblet cells are the most abundant epithelial cell type in the zebrafish intestinal epithelium. (Wallace et al. 2005; Ng et al. 2005).

To facilitate studying the digestive system in zebrafish, Field et al. (2003) established a transgenic strain of zebrafish, gutGFP, in which GFP is expressed in the zebrafish digestive system and the expression is restricted to the endoderm and endoderm-derived organs from 22 hours post fertilization (hpf) to adulthood. This transgenic line already assisted in studying liver morphogenesis in zebrafish (Field et al.

2003). Others utilized the gutGFP strain to characterize gut morphogenesis by studying the developmental stages that form the zebrafish intestinal epithelium over the first 5 days of development. They also used a different transgenic line, (Tg [nkx2.2a: mEGFP]), to study intestinal enteroendocrine cell development. In addition to EGFP expression in the brain, ventral neural tube, and developing pancreas, EGFP is also expressed in a subset of cells that display the typical morphology of enteroendocrine cells throughout the intestinal epithelium (Ng et al. 2005).

Her et al. (2004) produced another transgenic line using the intestinal fatty acid binding protein (ifabp) promoter to drive GFP expression in the developing intestine.

They showed that two GATA-type binding sites, one C/EBP binding site, and a novel

15-bp element within the I-FABP gene promoter contribute to intestine-specific gene expression and are functionally conserved between mammals and zebrafish. The outcome of this study recommended that zebrafish should be considered a suitable model for studies of gut development (Her et al. 2004).

Monoclonal antibodies were produced to recognize the absorptive and secretory cells in zebrafish gut epithelium, and the role of Delta-Notch signaling in generating different intestinal cell types were examined by Crosnier et al. 2005. BrdU labeling revealed a similarity between the renewing of gut epithelium in zebrafish and that of mammals. It was also found that there were several Notch receptors and ligands expressed in the gut, in particular, the Notch ligand DeltaD, which was expressed in cells of the secretory lineage. It was concluded that Delta-Notch signaling is necessary to produce a stable mixture of absorptive and secretory cells. These findings revealed the fundamental role of Notch signaling in the gut stem cell system and suggested that zebrafish are a useful model to study gut development, as well as the mechanisms regulating the regeneration of gut epithelium (Crosnier et al. 2005).

The zebrafish liver consists of three lobes that lie along the intestinal tract and is responsible for different vital functions such as bile production, blood detoxification, and the production of critical plasma proteins and clotting factors. Hepatocytes are considered the key component of the liver that performs most of the liver’s main functions in zebrafish (Le Douarin 1975; Cascio and Zaret 1991; Tao and Peng 2009).

The zebrafish liver does not have Kuppfer cells, its hepatocytes are not clearly structured in cords, and the typical portal triads are not obvious like in the liver of mammals (Van Der Ven et al. 2003).

The zebrafish liver has unique histological characteristics compared with that in mammals. Its portal veins, hepatic arteries, and large biliary ducts are distributed randomly within the hepatic parenchyma but are not grouped in portal tracts as in mammals. Furthermore, hepatocytes are arranged as tubules that enclose small bile

ducts rather than as bilayered hepatocyte plates as in mammals (Pack et al. 1996; Lorent et al. 2004). Zebrafish have some unique advantages for studying liver development.

Blood in zebrafish forms in the intermediate cell mass (ICM) first and subsequently in the posterior blood land (PBI) and finally in the kidney, but not in the liver (Detrich et al. 1995; Thisse and Zon 2002; Jin et al. 2009). Further, as zebrafish have a yolk that supplies it with nutrients during embryogenesis, zebrafish can remain alive and develop fairly normally for a few days without the cardiovascular system (Stainier 2001). These advantages allow the studies on liver development and disease in zebrafish even with mutations affecting blood development.

Field et al. (2003) took advantage of the zebrafish transparency, in conjunction with the use of gutGFP transgenic line, to study and analyze the liver development. They revealed two phases of liver morphogenesis: budding and growth. The budding phase begins when hepatocytes first aggregate, shortly after 24 hpf, and finishes with the formation of a hepatic duct at 50 hpf. The growth phase then begins, as evident by clear changes in the size and shape of the liver. The direction of liver budding was also found to be independent of the direction of intestinal looping. Furthermore, they discovered that endothelial cells in zebrafish are not required for liver budding (Field et al. 2003).

To further study liver development and its regeneration ability in zebrafish, Curado and co-authors recognized a mutation in the gene encoding translocase of the outer mitochondrial membrane 22 (Tomm22) that responsible for oliver mutant phenotype.

This mutation displays liver-specific defects which mainly affect the hepatocytes after their differentiation. Through employing morpholino antisense oligonucleotides (MOs) to knockdown Tomm22 levels in zebrafish they showed the vital role of Tomm22 in

hepatocytes persistence, and they detected the regeneration capability of zebrafish liver which they used it as an alternative model of liver regeneration to identify the role of

Wnt2b and fibroblast growth factor (Fgf) signaling in hepatocyte regeneration (Curado et al. 2010)

Recently, Li et al. (2012) investigated the role of Augmenter of Liver Regeneration

(ALR) protein, which acts as a hepatotrophic growth factor that stimulates hepatocyte proliferation and promotes liver regeneration after liver damage or partial hepatectomy in mammals, during vertebrate hepatogenesis. Zebrafish were also used as a model to study the function of augmenter of alr gene in liver organogenesis and this was performed by knockdown, with morpholino and via overexpression. It was demonstrated that alr is expressed in the zebrafish liver during hepatogenesis and deplete ALR protein suppress zebrafish liver development while overexpression of ALR induces liver growth (Li et al. 2012).

The zebrafish kidney is located in a retroperitoneal position, just ventral of the vertebral column. It has different head and trunk regions and has two nephrons with a glomerulus, proximal tubules, distal tubules, and collecting ducts resembling a mammal’s kidney (Drummond et al. 1998). Each nephron is partitioned into a glomerular vascular pole and tubular segments. The pronephric kidney tubules are organized into proximal and distal tubular regions, similar to mammals. However, the zebrafish does not contain a Loop of Henle (Wingert et al. 2007; Wingert and Davidson

2008).

Zebrafish nephron development and function have been investigated through studying the mechanisms that control nephron segmentation, in which each segment is

responsible for the secretion and reabsorption of particular solutes. Exploring and understanding nephron segmentation mechanisms may help understand many human renal diseases. Consequently, several studies have been conducted on zebrafish to resolve the enigma of segmentation mechanisms. Studies on embryonic kidney development in zebrafish have shown that pronephric nephrons segmentation in zebrafish resembling that of mammals. The outcomes of these investigations show that the zebrafish is the suitable model to examine the conserved developmental pathways that regulate nephron segmentation in both lower vertebrates and mammals (Wingert and Davidson 2008).

1.6.3 Zebrafish as a disease model

Researchers employed zebrafish as a model to studying human diseases such as genetic diseases, kidney development and disease, cancer, cardiovascular disorders, angiogenesis, neurological diseases, liver disease, immunological studies and

Alzheimer's disease. The transgenic technologies which stimulate tissue-specific expression of fluorescent proteins in a conjugation with advanced imaging microscopy, such as live imaging with single- or multi-photon confocal microscopy, has increased interests of scientists and promote the study of different pathologies using zebrafish embryos. With an increased understanding of the cellular mechanisms responsible for disease, we can use the knowledge obtained from the zebrafish for the development of therapeutics.

The development and function of zebrafish organs are strikingly similar to those of humans and the ease of originating mutant or transgenic fish has assisted the generation of disease models. Zebrafish share a high genetic similarity to humans and

approximately 70% of all human disease genes have functional homologs in zebrafish

(Langheinrich 2003; Bibhas Kar 2013) On the other hand, there are certain disadvantages in using zebrafish as models for human disease; zebrafish do not have some mammalian organs, such as the mammary glands, and phenotypic characteristics of diseases caused by orthologous genes can be very different in fish and humans. In addition, the zebrafish genome contains many duplicate genes, which may result in the ancestral functions being divided between the two gene copies (subfunctionalization) or acquire new functions (neofunctionalization) (Postlethwait et al. 2004).

Polycystic kidney disease (PKD), a common human genetic disease, is the result of complete enlargement of kidney tubule lumens due to over-production of epithelial cells leading to kidney fibrosis and renal failure when the cysts exist in sufficient size and number (Calvet and Grantham 2001). Indeed, a relatively large number of recessive mutations have been identified that are associated with cystic pronephroi and affect the development of the pronephros in zebrafish. Apparent glomerular or tubular cystic formation is the main feature of most of these mutations. It was observed that these mutants span the principal stages of nephrogenesis, such as nephron patterning and the development of epithelial polarity (Drummond et al. 1998). Others have discovered 12 genes that, when mutated, can cause pronephric cysts in zebrafish and two of these genes have already been connected to human cystic kidney diseases (Sun et al. 2004).

These observations confirmed the concept that preservation of lumen size and epithelial cell shape is a complicated process which is regulated by several cellular proteins or signaling pathways as a relatively large number of genes are required to maintain tubule structure (Drummond 2005).

To investigate inflammatory bowel disease (IBD), adult zebrafish was used to investigate colitis disease. Treating zebrafish with oxazolone generates an intestinal epithelial deterioration and goblet cell depletion in treated fish. This was accompanied by an increase in il1b, tnf, and il10 gene production, in addition to eosinophil infiltration into the zebrafish intestine. The only disadvantage of using adult zebrafish in such studies is its non-transparency, which critically restricts imaging (Brugman et al, 2009).

The MPO reporter zebrafish strain (Tg(BACmpx:GFP)i114) was used to investigate IBD, and larval zebrafish model of colitis was developed by administrating 3 dpf zebrafish to hapten 2,4,6-trinitrobenzene sulfonic acid (TNBS), which is a common mammalian model of colitis that utilized to study acute and chronic intestinal inflammation (Oehlers et al. 2011; Fleming et al. 2010). Fluorescent imaging of live zebrafish larvae were used to examine the histological structure of the gut. Typical changes in gut architecture were recognized all over the zebrafish gut, which may be linked to colitis, in addition to serious changes in goblet cell number. It has been concluded that zebrafish larvae are an excellent model of gut physiology and pathology, which related to human disease. Oehlers et al. (2011) observed an increase in gut- specific il1b expression, intestinal lipid metabolism alteration, goblet cell hypertrophy, and intestinal shortening. On the other hand, they did not detect any histological impairment in the intestinal epithelial cell and proposed a lack of any serious intestinal injury (Oehlers et al. 2011).

Recently, Goldsmith et al. (2013) adopted a model of epithelial injury in zebrafish, by administering 5 dpf zebrafish NSAID glafenine for up to 24 hours, which is a non- steroidal anti-inflammatory drug to investigate the role of μ-opioid receptor (MOR)

signaling on the unfolded protein response (UPR), which is a cellular stress response associated to the endoplasmic reticulum. This study was conducted by utilizing transmission electron microscope in conjugation with acridine orange dye. The authors observed a significant increase in intestinal epithelial cell apoptosis in the lumen with disrupted intestinal structure and arrested cell stress responses. The intestinal epithelial cells showed indications of endoplasmic reticulum (ER) and mitochondrial stress followed by reduction of the unfolded protein response (UPR) coupled to an inadequate activation of downstream UPR mediators such atf6 and s-xbp1. The study findings revealed that [D-Arg2,Lys4]-dermorphin-(1–4)-amide (DALDA) hinders glafenine-induced epithelial injury through promoting operational UPR (Goldsmith et al. 2013).

Fatty liver disease is considered one of most common disease in western nations.

The first stage of nonalcoholic fatty liver disease (NAFLD) is the aggregation of lipid droplets in hepatocytes, which is called steatosis. This disease can develop from steatosis to hepatocellular injury, fibrosis, cirrhosis, and end with liver failure in human.

Hepatocytes in NAFLD may utilized lipid droplets to provide an adequate energy resource, which may lead to extra hepatocyte dysfunction (Mantena et al. 2008;

Breitkopf et al. 2009). Therefore, significant efforts have been devoted to developing effective models of zebrafish larvae to study liver pathogenesis that could be used in drug discovery. One of the reasons to choose zebrafish is that several studies have revealed that hepatobiliary disease in zebrafish larvae resembles that in mammals (Chu and Sadler 2009; Schlegel 2012). Also, toxins can be easily administrated to zebrafish, which may cause fatty liver disease in humans such as ethanol (Howarth et al. 2011) or

tunicamycin (Tm) (Howarth et al. 2013). It was found that not all hepatocyte functions were disrupted in both models, which treated with ethanol and tunicamycin, while some features were only affected in response to EtOH. They also applied ethanol and tunicamycin to detect whether lipids in hepatocytes inducing hepatic dysfunction in

ALD or not. These findings illustrate the efficacy of zebrafish larvae as a disease model for studying fatty liver disease which generally done in mammals.

Autosomal dominant polycystic liver disease (ADPLD), which is a rare progressive disorder of the biliary epithelium, caused by mutations in one of two genes, sec63 or prkcsh and ends with enlargement of multiple cysts in the liver (Strazzabosco and

Somlo 2011). To investigate hepatic cystogenesis in (ADPLD), Tietz Bogert et al.

(2013) established zebrafish models by inducing mutations in the sec63 and prkcsh genes that lead to hepatic cystogenesis in humans. These zebrafish models were also used to test the effects of several therapeutic agents on hepatic cyst growth. Mutations in sec63 or prkcsh affected a particular set of proteins that interfere with cholangiocyte growth, proliferation, and secretion, by which contributing to cystogenesis. It was showed that the zebrafish is a successful model in representing polycystic liver disease

(PLD) with characteristic hepatic and pathogenic features, which is similar to human liver pathology. The authors also concluded that this model is helpful in investigating the mechanisms of cyst formation and progressive growth, and in evaluating alternative treatments of inherited cystic diseases (Tietz Bogert et al. 2013).

1.7 Aims of this study

Different components of the plasma membrane have pivotal implications in organizing and maintaining the structure and function of polarized epithelial cells. One of these important players is lipid rafts, which contain a high level of cholesterol and sphingolipids. It has been hypothesized that these microdomains play a significant role in many cell functions such as cell signaling and membrane trafficking. Apical membranes possess different compositions and functions from that of basolateral membranes, and it has been suggested that the localization of lipid rafts is higher at apical membranes than basolateral membranes, which may contribute to the function of polarized epithelial cells.

Polarity proteins, which preferentially localized at apical or basolateral membranes, also play a central role in organizing the structure and function of polarized epithelial cells including migration, differentiation, cell shape, and cell division. Polarity proteins have also significant implications in cell morphogenesis and polarized protein trafficking.

The composition and structure of plasma membranes is critical for cell function, but it has been challenging to study the organization of cellular membranes directly in vivo for various reasons. Most of the lipid rafts studies were performed on model membrane, or in cells in culture using various techniques, include biochemical approaches and imaging methods. A better understanding of the cell membrane and its organization can be achieved by visualizing cells in an intact vertebrate organism.

To date, lipid rafts, polarity proteins, and their role in cell structure and function have mainly been studied separately so that the relationship between membrane order 63

and polarity proteins and how membrane domains are regulated by the cell has remained unexplored. This study proposes the hypothesis that there is a correlation between membrane order and the localization of polarity proteins in polarized epithelial cells. Therefore, this study aims to address four questions:

1. Is membrane order of polarized epithelial cells changing during zebrafish

development stages (Chapter 3)?

To address this question, zebrafish larvae were stained with Laurdan at different development stages, namely at 3, 4, 6, 8 and 11 dpf. Membrane order was measured in the gut, kidney, and liver of live and fixed zebrafish larvae by 2-photon (2P) microscopy.

2. Can membrane order be manipulated by adding and depleting sterols in polarized

epithelial cells (Chapter 4)?

Here, zebrafish larvae at 6 dpf were treated with 7KC, cholesterol-mβCD, and mβCD, stained with laurdan, and membrane order measured in gut, kidney, and liver.

Another group of zebrafish larvae were treated with 7KC, mβCD, cholesterol-mβCD, and mβCD at 4 dpf then allowed to develop till 6 dpf, at which stage GP values were measured in gut, kidney, and liver. Immunofluorescence was performed on untreated and sterol-treated zebrafish to examine the tissue architecture and quantify the apical localization of aPKC in polarized epithelial cells.

3. Does polarity protein expression regulate membrane ordered in polarized epithelial

cells (Chapter 5)?

In this part of the study, zebrafish larvae were injected with morpholino to knockdown Par3 and Crb3a polarity proteins. Zebrafish were stained with Laurdan at 4 dpf and membrane order was measured in gut, kidney, and liver of knockdown and un-

injected, control larvae. Immunofluorescence with anti-aPKC antibody and anti-ZO-1 antibody was performed to analyze aPKC localization in apical and cytoplasmic region.

4. Does knock-down of flotillin proteins affect the membrane order and the apical

localization of polarity proteins in polarized epithelial cells (Chapter 6)?

Here, morpholinos were used to knockdown the lipid rafts proteins, Flot-1 and Flot-

2 and Laurdan microscopy and immunofluorescence used to assess membrane order and apical localization of polarity proteins.

In summary, this thesis presents how manipulating membrane order (Chapter 4) and limiting the expression of polarity proteins (Chapter 5) affects the localization of polarity proteins and vice versa, how the reduced expression of rafts proteins affects membrane order (Chapter 6). Taken together, the data establish a relationship between membrane order and the apical targeting of polarity proteins in gut, kidney, and liver of developing zebrafish larvae.

Chapter 2: Materials and Methods

2.1 Zebrafish

2.1.1 Zebrafish strain and husbandry

The Casper strain of zebrafish was used because of their transparent phenotype during all their developmental stages. Zebrafish were raised and maintained in the Brain and Mind Research Institute (BMRI) fish facility (University of Sydney), at pH (7.4), temperature (28°C) and salinity controlled conditions. Zebrafish were maintained on a regular light-dark cycle with 14 hours of light and 10 hours of dark.

2.1.2 Set up mating, collecting and sorting zebrafish embryos

In the afternoon or early evening of the day prior to embryo collection, fish were set up for timed mating in a small breeding tank consisting of a base tank, removable inner container (sieve) that holds the fish, and a plastic lid. Using a small net, two males, and three females were transferred to the breeding tank and the tank covered with a plastic lid. The following day, egg production was initiated following commencement of the light cycle. To collect the zebrafish eggs, the sieve tank containing the fish was transferred to another base tank. The water from the mating tank was then poured through a plastic tea strainer to collect the eggs. Using a squeeze bottle filled with E3 medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, 0.00001%

(w/v) methylene blue), the eggs were rinsed from the tea strainer into a Petri dish, and the Petri dish labelled with same tank data. After 24 hours, using a dissecting microscope, approximately 60 embryos that developed normally were sorted from the dish using a flexible plastic pipette and transferred to a new Petri dish filled with E3 66

medium and incubated at 28°C for embryo development (Nüsslein-Volhard and Dahm

2002).

2.2 Laurdan microscopy

2.2.1 Laurdan staining and mounting

A 5 mM Laurdan (Invitrogen, D250) solution was prepared in dimethyl sulfoxide

(DMSO) (Sigma, D2650). Zebraﬁsh larvae were transferred with 1 ml of E3 medium to a small Petri dish or multi-well tissue culture plate. 10 µl of 5 mM Laurdan solution was then added to the 1ml of E3 medium to reach a final Laurdan concentration of 50 µM.

Larvae were stained for 30 min before washing with PBS, followed by a 30 min recovery period in E3 medium. To anesthetize and mount the larvae, larvae were transferred to an eppendorf tube containing E3 medium. The E3 medium was then sucked up by plastic pipette and 50 µl of 0.04% ethyl 3-aminobenzoate methanesulfonate (Tricaine) (Sigma, E10521) and 1 ml of 1% low melting agarose

(Sigma, A9414) were added and mixed. The whole mixture was then transferred to a small Petri dish and left to solidify at room temperature for imaging.

2.2.2 Imaging and image analysis

An inverted confocal laser-scanning epi-fluorescence microscope (SP5; Leica

Microsystems, Wetzlar, Germany) was used for image acquisition with excitation at 800 nm using a femtosecond-pulsed Ti: Sapphire laser (Mai-Tai; Spectra-Physics, Mountain

View, CA). The wavelengths 400–460 nm and 470-530 nm were used for fluorescence detection using internal photomultiplier tubes (PMT) with the confocal pinhole open and recorded as 8-bit, 512 X 512 pixels digital images. Scan speed was set at 400 Hz; 67

and the line average to 6. Zebrafish larvae dishes were placed on a 63× 1.3 NA glycerol- immersion objective lens. All acquired ordered (ch00) and disordered (ch01) images were saved as TIFF files for further analysis to quantify GP values. To compensate for variations in PMT settings and sensitivity, a solution of 0.5 µM Laurdan in DMSO was imaged at the end of each imaging session; see the ‘G factor correction’ in Section 2.2.3.

These were then used to construct GP images using ImageJ (National Institutes of

Health, Bethesda, MD). The GP images were pseudo-colored, which were merged with mean fluorescence intensity images, to maintain structural information, using the hue- saturation-brightness (HSB) color space.

2.2.3 Calibration factor (G-factor) calculation

Because the GP function is based on the relative intensity of the blue and green channels, acquisition parameters such as objective lens, PMT gain and offset critically influence the relative intensity in the two channels and hence the final GP values.

Therefore, it is essential to calibrate the relative intensity of the two channels to obtain an absolute measurement of the GP values. The calibration is use to correct the GP values by introducing the G-factor into the GP equation applied that is used to calculate the GP images. The factor G serves to calibrate the relative intensity of the two channels.

Here, I used intensity images of two spectral channels and calculated a normalized ratio according to the following formula.

Where I is the intensity in each pixel in the image acquired in the respective spectral channel and G is the calibration factor.

To determine the G factor, the differences in PMT settings and sensitivity were compensated through calibrating image acquisition, and this was accomplished by adjusting the instrument settings identical to those used for the experiment

(objective lens, immersion media, PMT gains, and offsets, zoom factor). A solution at

~1:100 dilution of 5 mM Laurdan in DMSO was imaged with three different laser powers (the same power as that used for imaging the sample, in addition to 50% higher power and 50% lower power). To calculate the G factor, the mean value of the ordered

(channel 1) and disordered (channel 2) intensity images were recorded and the G factor was calculated according to:

The theoretical reference value for Laurdan in DMSO, GPRef, is 0.207 at 22℃.

This was repeated for all G-factor images acquired at different laser powers and the resulting G factors were averaged. The resulted value of the G factor were applied to analyze and quantity the GP value of the region of interest (ROI) in each image.

2.2.4 Image analysis and data acquisition

In order to quantify membrane order of the epithelial cells, 10 GP images were acquired from 10 larvae at 3 dpf stage. GP images were generated from blue and green intensity images then pseudo-colored with highly ordered membrane regions represented in purple-red colors and lower order domains represented in orange-green while the lowest membrane order regions represented in green-blue colors (Figure 2-1). 69

Actually, any pixels with low intensity in blue and green channels need to be excluded; this was done by thresholding the raw input images. This also guarantees that the quantification includes only stained membranes with a decent signal•to•noise ratio.

The grayscale GP image was generated from the two intensity images that collect

Laurdan emission between 400-460 nm and 470-530 nm. To display the image data, the grayscale image was pseudo•colored by using a rainbow RGB (blue•red) lookup table

(LUT) as an example, and then thresholded pixels were colored black.

Five ROI were selected from each of the gut, kidney, and liver GP images by select regions at the apex, the lateral and the inner of the epithelial cell, which represent the apical, basolateral and intracellular regions respectively (Figure 3-4, 5 and 6).

Membrane GP measurements were extracted from the representative grayscale GP images and were calculated from the mean intensity values for that ROI of 3 dpf larvae images. Grayscale GP images were used to extract GP values of membranes of interest.

To generate images, grayscale GP images were pseudo-colored and then merged with the mean intensity images to generate hue•saturation-brightness (HSB) color space. The pseudocolored GP image merged with the intensity image by multiplying each color component by the intensity value in that pixel. (Figure 2-1). Then the (0:255) digital number (DN) values were converted into − 1: + 1 GP values by using this formula GP =

(Owen et al. 2012).

Figure 2- 1: Generating GP images and pseudo-colored images for data acquisition. The GP image is generated from the two intensity images that collect Laurdan emission between 400-460 nm (shown in green) and 470-530 nm (shown in red). Grayscale GP images were used to extract GP values of membranes of interested or pseudo-colored. The pseudo-colored image were merged with the mean intensity images to generate hue•saturation•brightness (HSB) color space. Pixels that contained only background Laurdan fluorescence were colored black. The color range of the lookup table (LUT) was set to -1 to + 1 with GP = + 1 in red representing ordered membranes and GP = − 1 in blue which representing fluid membranes.

2.3 Immunofluorescence staining and confocal imaging

2.3.1 Antibody staining of whole mounted zebrafish

Larvae were fixed in 4% paraformaldehyde (PFA) overnight at 4°C. The following day they were washed in PBST (PBS pH7.4 + 0.1% Tween20 (Sigma, P1379)) for 5 min followed by distilled H2O for 5 min. Larvae were then frozen in pre-chilled acetone at -20°C for 7 min to permeabilize the tissue. Chilled larvae were again washed in distilled H2O for 5 min followed by washing in PBST buffer for 5 min. Larvae were then incubated with 2% normal goat serum (NGS) (Sigma, G9023) in PBDT (PBS

pH7.4 + 1% bovine serum albumin (BSA) (Sigma, A9056) + 1% DMSO + 0.1%

Tween-20) for 1 hour at room temperature to block non-specific binding sites. Larvae were then incubated with an antibody cocktail containing primary mouse monoclonal antibody ZO-1 (1:300) (Invitrogen, 339100), wheat germ agglutinin Alexa Fluor 594

Conjugate (WGA) (1:50) (Invitrogen, W11262), and Cholera toxin subunit B Alexa

Fluor 647 Conjugate (CTxB) (Invitrogen, C-34778) in 2% NGS-PBDT overnight at 4°C with gentle rocking. The following day, larvae were washed eight times for 15 min each in PBST at room temperature. Following washing, larvae were treated with Alexa Fluor

488 goat anti-mouse IgG secondary antibody (Invitrogen, A11001) diluted 1:500 in 2%

NGS-PBDT overnight at 4°C with gentle rocking. The following day, larvae were washed eight times for 15 min each in PBST, then larvae treated with DAPI (0.1%

DAPI in 50 ml PBS; Invitrogen, D1306) for 30 min to stain the nuclei. After that, larvae were mounted with Dako mounting media (Dako, S302380) and then imaged.

2.3.2 Confocal imaging of zebrafish

Imaging was conducted on an inverted confocal laser-scanning microscope (SP5;

Leica Microsystems, Wetzlar, Germany) with excitation at 800 nm. Fluorescence was detected in the ranges 380-460 nm for DAPI), 510-560 nm for ZO-1, 597-620 nm for

WGA and 640-700 nm for CTxB using internal photomultiplier tubes (PMT) with the pinhole size set to 1 Airy. The 63x 1.3 NA glycerol-immersion objective lens was selected for imaging.

2.3.3 Antibody staining of sectioned zebrafish

2.3.3.1 Zebrafish sectioning using a vibratome

Larvae were fixed with 4% PFA overnight at 4°C and the following day washed in PBS three times for 5 min each. 4% low melting point agarose was prepared in

PBS and stored at 55°C in a water bath until needed. Agarose was then poured into plastic sectioning molds and the larvae added at a lateral orientation. The agarose block was kept at room temperature until set and then removed from the mold using a spatula. The bottom end of the agarose block was sliced with a razor blade to create an even surface and then glued to the vibratome stage, which then attached to the vibratome (Vibratome, Inc.). The parameters of the vibratome were set at a speed of

3 and a thickness of 50 μm). Once the larvae were sectioned, the desired sections were placed into a PBS-filled multi-well tissue culture dish.

2.3.3.2 Antibody labelling

In a multi-well tissue culture dish zebrafish sections were incubated with 2% NGS-

PBDT for 3 hours at room temperature to block non-specific bindings. Without washing, sections were then incubated with an antibody cocktail containing primary mouse monoclonal antibody ZO-1 (1:300) and rabbit anti-PKC ζ antibody (1:500) (

Santa Cruz Biotechnology (C-20): sc-216) in 2% NGS-PBDT overnight at 4°C with gentle rocking. The following day, sections were washed four times (10 min, 15 min, 30 min, and 60 min, each) in PBT (1% BSA + PBS + 0. 1% Tween-20). Sections were then incubated with Alexa Fluor 488 goat anti-mouse IgG secondary antibody and Alexa

Fluor 594 goat anti-rabbit IgG secondary antibody (Invitrogen, A-11012) diluted at

1:500 in 2% NGS-PBDT overnight at 4°C with gentle rocking. The following day sections were washed four times (10 min, 15 min, 30 min, and 60 min, each) in PBT.

Finally, sections were treated with DAPI (0.1% DAPI in 50 ml PBS) for 30 min to stain the nuclei. Sections were then washed three times in PBS for 5 min each and mounted on a slide using Dako mounting media as previously described (Jayachandran et al.

2010; Shim 2011).

2.3.4 Confocal imaging

Imaging was performed with an inverted confocal laser-scanning microscope (SP5;

Leica Microsystems, Wetzelar, Germany) with excitation at 800 nm. Fluorescence was detected in the ranges 380-460 nm for DAPI, 510-560 nm for ZO-1, and 597-620 nm for aPKC using an internal photomultiplier (PMT).

2.4 Preparation and quantification of sterol-cyclodextrin complexes 2.4.1 Preparation of Sterol-cyclodextrin complexes

Separate 15 mg/ml cholesterol and 7KC solutions in ethanol were prepared

(Cholesterol = 5 cholesten-3β-ol, Sigma, C8667) (7KC = 5-cholesten-3β-OL-7-ONE,

Steraloids, C6970-000). A 5% mβCD stock was prepared by dissolving 500 mg of mβCD in 10 ml distilled H2O (mβCD = methyl-β-cyclodextrin, Sigma, C4555). A hotplate stirrer containing a heat block was set to 80°C and HPLC screw-top vials containing tiny stirring fleas inserted. 400 µl of 5% mβCD stock was then added to each vial and whilst stirring four 10 µl aliquots of either 15 mg/ml cholesterol or 7KC added to each vial; leaving 5-10 min between each 10 µl addition. After the fourth aliquot had been added, the preparations were left on the hotplate at 80°C for 1 hour uncovered. 74

After 1 hour, all aliquots of the prepared cholesterol-mβCD complexes were collected into a 15 ml falcon tube. All aliquots of the prepared 7KC-mβCD complexes were collected into another 15 ml falcon tube. Cholesterol-mβCD and 7KC-mβCD complexes were then frozen at -20°C until HPLC analysis (Klein et al. 1995).

2.4.2 Quantification of sterol-cyclodextrin complexes

The cholesterol-mβCD and 7KC-mβCD complexes were thawed at room temperature. 15 µl of cholesterol-mβCD and 7KC-mβCD complex were then added to separate 1.5 ml eppendorf tubes and made up to 1 ml with PBS; both the cholesterol and

7KC analysis were carried out in triplicate. 10 µl of 200 µM butylated hydroxytoluene

(BHT) (2, 6-Di-tert-butyl-4-methylphenol;Sigma, B1378) prepared in ethanol), 100 µl of 200 mM ethylenediaminetetraacetic acid disodium salt dehydrate (EDTA; Sigma,

ED2SS) prepared in distilled water and 500 μl of diluted sterol complex were added to a Kimax tube and made up to 1 ml with PBS. 2.5 ml of methanol and 5 ml of hexane were then added to the Kimax tube and the mixture vortexed for 15 seconds before centrifuging at 2500 rpm, 10°C for 15 min.

Following centrifugation, 4 ml of hexane (top layer) was removed and added to a

Kimble tube for evaporation to dryness in a speed vac concentrator (Thermo Scientific).

Samples were then reconstituted in 200 µl of mobile phase (MP210) (70% isopropanol and 30% acetonitrile) and transferred to HPLC vials for analysis. To determine peak retention time, a cholesterol / 7KC mixed standard was prepared by adding 10 µl of 15 mg/ml cholesterol and 10 µl of 15 mg/ml 7KC to a Kimble tube. This was then evaporated to dryness in a speed vac concentrator reconstituted in 180 µl of mobile phase MP210 and transferred to an HPLC vial for analysis.

The concentration of cholesterol and 7KC within each sterol-cyclodextrin complex was measured by HPLC using a Supelco reverse-phase C18 column (0.46×25 cm, 2 cm

Pelliguard column, 5 µm particle size; Sigma-Aldrich) run at room temperature with a flow rate of 1 ml/min using the mobile phase acetonitrile/isopropanol/water (44:54:2 v/v). Cholesterol and 7KC were detected using a UV detector at a wavelength of 210 nm and 234 nm, respectively. To quantify the concentration of sterol within each sterol- mßCD complex, the peak area of the eluted sterol was compared with a previously determined response factor for that sterol. The concentration of 7KC-mβCD was 4.97 mM, and cholesterol-mβCD was 5.64 mM.

2.5 Morpholino knockdown experiment

2.5.1 Morpholino Oligonucleotides (MOs) sequence

Four antisense morpholino oligonucleotides were synthesized by GeneTools LLC

(Cowallis, OR, USA), which were designed based on the 5’ UTR sequence to reduce protein expression of Par3, Crb3a, Flotillin-1a (Flot-1a), and Flotillin-2a (Flot-2a) in zebrafish larvae. Morpholinos sequences were as followed: par3 MO: 5’

TCAAAGGCTCCCGTGCTCTGGTGTC 3’(Wei et al. 2004), crb3a MO: 5'

AGCCCAACCTGCTGGATCATTTCCG 3' (Omori and Malicki 2006), flot-1a MO: 5’

TTTTAGACGTTGGCTGAATGATTAG 3' and flot-2a MO: 5’

CATTTTCCTTTCTGGACGCCTTTAA 3'.

2.5.2 Microinjection of morpholino into zebrafish embryos

2.5.2.1 Micropipette and microinjection chamber plate preparation

Micropipettes were prepared by heating and pulling borosilicate glass capillary tubes in a micropipette puller device (P-97 Flaming/Brown Micropipette Puller, Sutter

Instrument, USA); a 1mm OD glass capillary tube was pulled into two needles. To prepare the microinjection chamber plate, 1.5% low melting agarose was poured into

Petri dish then the mold plate, which designed into a wedge-shape to hold the embryos during injection, placed on the top of agarose. Microinjection chamber plates were then left to solidify at room temperature before being stored at 4ºC until use.

2.5.2.2 Morpholino loading

The distal tip of a micropipette was cut with a surgical razorblade to make an opening that was observable under a dissecting microscope at 50X magnification. The needle was loaded with 3 μl of injection material using a microloader pipette.

2.5.2.3 Injection volume calibration

Under the dissection microscope, the injection volume was tested by placing a drop of working solution (morpholino), using the micro-injector, onto micrometer oil.

The diameter of the drop was measured as it floated as a sphere on top of the oil. The duration and pressure of injection was then adjusted to 24 mms and 20 psi.

2.5.2.4 Zebrafish embryo preparation for microinjection

Zebrafish were set up for mating as described in Section 2.1.2. The eggs were collected at the 1-cell stage from the breeding tanks and rinsed with E3 medium in a

Petri dish. Unfertilized eggs and debris were removed. Embryos were arranged along the wedge-shaped troughs in the microinjection chamber plate by using a 3 ml transfer pipet. Some E3 media was then removed so that the embryos were shallowly submerged but not flooded.

2.5.2.5 Morpholino Oligonucleotides preparation

A 300 nmol morpholino oligonucleotide was specifically designed for a gene of interest from Gene Tools, LLC, and 300 µl of sterile water added to the lyophilized stock to make a 1 mM stock solution. The solution was then aliquotted and stored at -

20°C until use. On the day of injection, the morpholino stock solution was heated at

65°C for 5 min, then snap cooled on ice followed by a brief spin. Morpholino oligonucleotides were re-suspended at 1-2 μg/μl in distilled water containing 0.1% phenol red before injection into Casper strain embryos at the 1-4 cell stage.

2.5.2.6 Zebrafish microinjection

Embryos at the 1-4 cell stage were manipulated with the micropipette so that the cytoplasm of the embryo was visible under the dissecting microscope. The morpholino working solution was injected into the yolk at 1 nl per embryo. par3 MO was injected at

125 μM, crb3a MO at 250 μM, flot-1a MO, and flot-2a MO at 500 μM.

The dosages were adjusted to optimize the phenotype-to-toxicity ratio. The injected eggs were kept in E3 medium at 28°C to develop further. Unfertilized eggs were discarded the next day to maintain the appropriate developing conditions for embryos.

At 4 dpf stage, the larvae were observed for the phenotypes of interest as previously described (Rosen et al. 2009; Yuan and Sun 2009).

2.6 Western Blot

2.6.1 Preparation of total lysates

Zebrafish larvae were anaesthetized using Tricaine, and the yolk removed manually from ~ 20 larvae in cold Ringer’s solution (1.7 g NaCl, 0.15 g KCl, 0.2 g NaHCO3 in

500 ml H2O pH7). Larvae were then washed with ice-cold PBS and centrifuged at 1500 rpm for 5 min, and the supernatant discarded. The larvae were then homogenized in T-

PER extraction reagent (Pierce Biotechnology Inc., Rockford, IL, 78510) containing 2x complete protease inhibitor tablets (Roche, 04693132001) (one tablet of protease inhibitor in 5 ml T-PER) on ice (~0.75 μl/embryo) and then centrifuged at 14,000 rpm for 15 min to remove debris. The supernatant was transferred to a new tube, and the debris discarded. Protein concentration of the yolk lysates was determined by Nanodrop spectrophotometer (Nanodrop 1000 Thermo Scientific) and following protein quantification, cell lysates diluted to equal protein concentrations with T-PER extraction buffer containing complete protease inhibitors.

2.6.2 Immunoblot

2.6.2.1 Preparing cell lysates and electrophoresis

13 µl of whole embryo lysate, 5 µl of NuPAGE® LDS Sample Buffer (4X)

(Invitrogen, NP0007) and 2 µl of NuPAGE® Reducing Agent (10X) (Invitrogen,

NP0004) were added to an eppendorff tube and heated at 70°C for 10 min before centrifuging. An equal amount of lysate protein (18-33 µg) was loaded and electrophoresed at 180 V for 50 min through a NUPAGE 10% BisTris polyacrylamide gel (Invitrogen, NP0315) in MOPS SDS Running Buffer (Invitrogen, NP0001). 79

Separated proteins were electrotransferred to an Immobilon-P PVDF membrane

(Millipore, IPVH00010) by wet transfer at 30 V for 1 hour in NuPage Transfer Buffer

(Invitrogen, NP0006-1). Transfer efficiency was assessed by Ponceau S staining (Sigma,

P 7170) as previously described (Vihtelic et al. 1999).

2.6.2.2 Polarity and rafts proteins detection

Membrane was blocked in blocking buffer (5% BSA + TBS (pH 7.5) + 0.05%

Tween-20) at room temperature for 1 hour. Then, rabbit anti-Par3 polyclonal antibody

(Millipore, 70-330), rabbit anti-Crb3 monoclonal antibody (Uppsala University-

Sweden), mouse anti-Flotillin-1 monoclonal antibody (BD Biosciences, 610821) and

rabbit anti-Flotillin-2 polyclonal antibody (Santa Cruz Biotechnology, (H-90): sc-

25507) were diluted 1:1000 in 5% (w/v) BSA in TBST, and incubated on the membrane

overnight at 4°C. The following day, the membrane was washed in TBST (two brief

rinses, one 15 min wash, and two 5 min washes). Antibody binding was then detected

by incubating with anti-rabbit IgG or anti-mouse antibody conjugated to HRP (BioRad,

170-6515) diluted 1:3000 in 5% (w/v) BSA in TBST for 1 hour at room temperature.

The membrane was then washed in TBST (two brief rinses, one 15 min wash, and four

5 min washes). After washing in TBST, the blot was rinsed in water and incubated with

Pierce Enhanced Chemiluminescence (ECL) Western Blotting Substrate (Thermo

Scientific, 32209). Phosphorescent bands were imaged in an ImageQuant LAS 4000

mini (GE Healthcare, 28-9558-13) with ‘super’ CCD sensitivity and incremental

exposures recorded every 10 seconds for up to two min.

2.6.2.3 Stripping the membrane

Following proteins detection, the membrane was incubated in stripping buffer

(Thermo Scientific, 46430) at room temperature on a rotating table for 20 min then stripping buffer was removed and membrane rinsed two times for 10 min each at room temperature in PBST. The membrane was re-blocked as in Section 2.6.2.1, and proceeded with antibody binding. The expression of β–actin (1:1000) was used as a control to confirm that an equivalent amount of protein extract was loaded in each lane.

2.7 Statistical Analyses

Statistical comparisons between two populations were made using Student’s t-test assuming unequal variances. Comparison between multiple groups was performed by one-way ANOVA with Bonferroni post-test, and results were considered statistically significant at P < 0.05. Unless stated otherwise, data were presented as mean ± standard deviation (SD) from at least three replicates of a single experiment. All statistical analyses were performed with Prism software (GraphPad Software Inc., USA).

Chapter 3: Membrane order of polarized epithelial cells during the development of zebrafish larvae

3.1 Introduction

Plasma membrane microdomains such as lipid rafts have received an enormous amount of interest due to their significance in various cellular functions including signaling cascades (Simons and Toomre 2000), protein and lipid sorting, trafficking

(Ikonen 2001), and cell adhesion and migration (Gómez-Moutón et al. 2001). The most challenging aspect of lipid rafts in terms of experimental characterization is their proposed small size and short lifespan, making it difficult to analyze, visualize and explore their structure and function.

Cell membranes are composed of a lipid bilayer which is composed of two opposite leaflets. The lipid bilayer is mainly held together by non-covalent forces (hydrophobic interactions) so that lipids can move laterally within the bilayer in a rapid and natural process called "lateral diffusion” to organize the cell functions. Two lipid phases exist in the bilayer membrane, the liquid-ordered phase (lo) and liquid-disordered phase (ld). It has been hypothesized that in the liquid-ordered phase, lipids are arranged on a two- dimensional triangular lattice in the plane of the membrane. In phase segregation, the membrane bilayer converts from a highly liquid-ordered phase to a disordered liquid phase and vice versa. Cholesterol and phospholipids in the bilayer have the ability to form a liquid-ordered (lo) phase that can coexist with a liquid-disordered phase (ld), the latter has a lower concentration of cholesterol (Simons and Ikonen 1997; Simons and

Meers 1988).

The techniques used to study lipid rafts include both biochemical methods and imaging methods. Biochemical techniques that rely on detergents such as Triton X-100,

Lubrol WX, and Brij 98 (Schuck et al. 2003) are considered invasive procedures. There are some concerns in using invasive methods as they may disrupt and re-organize lipid and protein complexes (Shogomori and Brown 2003). It is uncertain whether the addition of detergents to cells will maintain membrane structures. Imaging techniques are an alternative to investigate membrane order in the cell membrane, where a membrane probe or marker can be readout with various fluorescence microscopy techniques (Bagatolli and Gratton 2000; Gaus et al. 2003; Owen et al. 2006). Many imaging methods have failed to confirm the existence of rafts in living cells. Meanwhile,

Owen and his group have provided evidence for lipid organization in the cell membrane.

Owen et al. 2006 used the di-4-ANEPPDHQ probe, which is an environmentally sensitive fluorescent probe for lipid membranes, with FLIM to visualize liquid-ordered and liquid-disordered phases in model membrane. They succeeded in applying this approach to live cells, and observed variable levels of ordered phase in live epithelial cells (Owen et al. 2006). Jin et al., 2006 also used di-4-ANEPPDHQ dye with both single photon fluorescence and two-photon fluorescence microscopy. They imaged the phase separation in giant unilamellar vesicles (GUVs) and di-4-ANEPPDHQ dye was able to differentiate liquid-ordered and liquid-disordered phases in those GUVs (Jin et al. 2006).

Here, I used Laurdan in conjunction with two-photon microscopy to quantify membrane order. The Laurdan dye integrates itself into the cell membranes and is only emit fluorescence in lipid environments (Bagatolli et al. 1998). Laurdan is an

environmentally sensitive probe, meaning that its emission spectra depend on the condition of the surrounding membrane. More specifically, Laurdan senses the degree of water within the lipid bilayer; when water is largely excluded, and lipids are tightly packed such as in liquid-ordered membranes, the emission peak of Laurdan is around

440 nm. In more fluid membranes, such as liquid-disordered membrane phases, water can partially penetrate between the lipid molecules in the bilayer and this increase

Laurdan’s emission peak to around 490 nm. A relative measurement of the membrane order, the Generalized Polarization (GP), can be calculated by measuring the intensity of Laurdan fluorescence at both emission regions at the same time, and calculating a normalized ratio between the two channels. Here, Laurdan fluorescence emission is recorded across the range of 400–460 nm for ordered membranes and 470–530 nm for fluid membranes.

The calculation of the Generalized Polarization returns a number between -1 (fluid) and +1 (ordered) which can be used in quantifying membrane order of the epithelial cells (Parasassi and Krasnowska 1998; Dietrich et al., 2001).

A better understanding membrane organization and fluidity can be achieved by visualizing and imaging cell membranes in a whole vertebrate organism rather than in model membranes or tissue culture models of epithelial cells. This is because in vivo studies retain the cellular arrangement within organs, which is critical for the polarization of epithelial cells. Hence it is expected that the physical-chemical properties of cell membrane are unperturbed in an in vivo model.

Here, zebrafish was chosen as a model to investigate membrane order of polarized epithelial cells. As a vertebrate model, zebrafish has many advantages over other animal

models such as mouse and rabbit. It is robust fish, breeds all year around, and produces large numbers of eggs with short development time. Further, raising zebrafish in the lab is safe and affordable. In addition, zebrafish embryos are transparent and consistent in their size, which make it easy to observe the cellular organization with light microscopy.

Zebrafish is characterized by its rapid development; by day 3 the larva completed most of its morphogenesis, and it continues to grow rapidly. The gut can be seen from two day onwards, and the yolk extension begins to vanish. The early larvae gradually start to perform different actions at this stage and start to swim, moving their jaws and eyes and later, performing rapid escape reactions, the seeking of prey, and feeding (Kimmel et al., 1995).

Zebrafish was the first whole living vertebrate organism, in which the membrane order of polarized epithelial cells was successfully imaged and quantified in a proof-of- principle study (Owen et al. 2010). This was accomplished by using the Laurdan dye in conjunction with multiphoton microscopy. It was shown that apical membrane of polarized epithelial cells had higher membrane order than basolateral membrane, which is consistent with the observation in cultured cells (Vieira et al. 2006). This study revealed a new way to quantify membrane order in live zebrafish, which unlocked the doors to explore membrane organization in vivo (Owen et al. 2010). Here, this approach was extended to quantify membrane order in polarized epithelial cells during development.

Transparency is one of interesting features in zebrafish, but unfortunately, pigmentation begins during embryogenesis and larvae lose their transparency gradually

(Kimmel et al. 1995). Pigmentation is one of the significant obstacles in imaging with

confocal microscopy because melanophores may generate significant amounts of fluorescence. To overcome these obstacles, different techniques were developed to produce transparent zebrafish. These techniques either generate transparent strains through genomic manipulation or by inhibiting melanogenesis by using chemical compounds such as 1-phenyl 2-thiourea (PTU) (Whittaker 1966; Westerfield 1995), which keeps embryos transparent as long as the PTU levels are sustained. However, through several studies, it has been found that using PTU may result in a high embryo mortality, reduced hatching frequency, and abnormal development without any structural malformations (teratogenesis) (Karlsson et al. 2001). For this reason, we selected a mutant strain called Casper to perform our study. This strain shows a complete absence of all melanocytes and iridophores during embryogenesis and adulthood stages, which make the fish almost totally transparent (Figure 3-1).

Casper showed better optical transparency compared to several commonly used zebrafish pigmentation mutants such as albino, rose, and panther. Some of the zebrafish organs like the heart, intestinal tube and liver, are more easily visible in Casper than other mutants. The Casper mutant produce large numbers of viable offspring at predictable mendelian ratios and with wild type heterozygous phenotype (White et al.

2008).

Wild type-AB Casper Casper

Figure 3-1: Wild type-AB zebrafish and mutant strain Casper. The photographs show the differences between wild type and mutant type Casper in adulthood stage. In contrast to wild type, Casper shows complete absence of melanocytes and iridophores during all its life, and its internal organs are visible by eye.

This chapter describes the characterization of membrane order in different tissues of live zebrafish such as gut, kidney, and liver, which possess highly polarized epithelial cells. I quantified membrane order (as GP values) of polarized epithelial cells of gut, kidney, and liver, during different developmental stages, 3, 4, 6, and 8 and 11 dpf (days’ post fertilization) in zebrafish larvae. The GP values between day 3, 4, 6, 8, and 11 were compared to identify differences that may occur in membrane order of apical, basolateral and intracellular membranes during zebrafish development stages.

3.2 Development of an imaging protocol for membrane order measurement in zebrafish larvae

To image and quantify membrane order of polarized epithelial cells of the gut, kidney, and liver from 3 dpf to 11 dpf, we raised zebrafish larvae and stained them with

Laurdan and then acquired and analyzed intensity images to quantify GP values. In order to validate the data, we first examined the effect of Laurdan on zebrafish development and tested the effect of paraformaldehyde (PFA) fixation on GP values measurements.

3.2.1 Laurdan staining of zebrafish larvae

Casper mutant strain was the appropriate choice to perform our investigation because of their transparency during the whole developmental stages that overcome problems with other transparent mutant strains (White et al. 2008). Zebrafish were mated and embryos collected and developed as described in Section 2.1.2 in Materials and Methods chapter.

To stain the epithelial cells of gut, kidney and liver of zebrafish larvae, a clutch of 3 dpf larvae were stained and imaged as described in Section 2.2 Materials and Methods chapter.

3.2.2 Effects of Laurdan on zebrafish development

To examine the effect of Laurdan on zebraﬁsh development, zebrafish were grown and mated, and embryos were collected. Three groups of embryos, with ∼20 embryos in each group, were raised till they became 3 dpf larvae and then transferred into small

Petri dish with 1 ml of E3 medium. At that point, 0 µl, 10 µl and 20 µl of 5 mM

Laurdan was added to 1 ml E3 medium of two groups of larvae to give a final concentration of 0 µM, 50 µM and 100 µM Laurdan, respectively, and incubated for an hour at 28°C. Larvae were washed three times with PBS and returned back to E3 medium at 28°C. At day 5, all three groups were transferred to the fish tank to start feeding, and all of the 3 groups kept alive at the aquarium to develop normally. After a month, all groups were examined for any abnormal development or behavior, and no adverse effects were detected on zebrafish development in both treated groups compared with the Control group (Figure 3-2).

Figure 3-2: Laurdan staining has no effect on zebrafish development. Two groups of zebrafish larvae were stained at 3 dpf with 50 µM and 100 µM Laurdan for 1 h, respectively. After a month, no adverse effects were observed on zebrafish development or behavior. Scale bar = 2 mm.

3.2.3 Image analysis and data acquisition

Larvae at 4 dpf, were stained, imaged live and GP values were recorded as described in Section 2.2 in Materials and Methods chapter. At 5 dpf, larvae were transferred to the feeding tank as they need to be fed at this stage and recovered to image 6 pdf, 8 dpf, and 11 dpf larvae, again live as described for larvae at 3 dpf and 4 dpf.

3.3 Results of membrane order measurements in developing zebrafish

3.3.1 Membrane order of epithelial cells in the gut during development

The zebrafish gut starts forming, from the gut primordium, around 42 hpf, to develop into a hollow tube (Ober et al. 2003). Around 3 dpf, the gut can be easily observed, and its epithelial cells were clearly seen in the GP images (Figure 3-4). At this stage, the gut of zebrafish larvae is a single layer of columnar-shaped epithelial cells that lay over a connective tissue, which is similar to the lamina propria of the mammalian small intestine. At the apex of these epithelial cells are apical membranes,

which face the lumen while the basolateral membranes are located at the lateral and bottom of these columnar-shaped epithelial cells while intracellular membranes are the interior of these cells (Figure 3-3).

Gut

IM Lumen AM

BM Connective tissue

Figure 3-3: GP image of zebrafish gut. Representative GP image of zebrafish gut show the columnar- shaped epithelial cells adjacent to connective tissue. Apical membrane (AM) (highlighted in white), basolateral membrane (BM) (indicated by the yellow selection) and intracellular membrane (IM) (highlighted in blue).

Figure 3-4: GP images of zebrafish gut, kidney, and liver in 3-11 dpf development stages. At 3-11 dpf, zebrafish larvae were stained with Laurdan, intensity images of the gut, kidney, and liver recorded with a 2-photon microscope and the images converted to GP images. GP images were pseudo-colored with red indicating ordered membranes with high GP values, orange-green indicated fluid membranes with moderate GP values and green-blue indicated more fluid membranes with low GP values. Apical (AM) and basolateral (BM) membranes were identified from the tissue morphology as shown in the zoomed regions (white squares). Scale bar = 20 μm. AM = apical membrane, BM = basolateral membrane and IM = intracellular membrane. ** gut lumen.

Figure 3-5: Comparison of GP values of apical (AM), basolateral (BM) and intracellular membranes (IM) in epithelial cells of the gut, kidney, and liver in 3-11 dpf zebrafish larvae. Each symbol represents one ROI obtained from 10 larvae with a total of 50 ROI at each development stage. **** indicates a statistical difference of P < 0.0001 (one-way ANOVA with Bonferroni post-test).

As it is shown in (Figure 3-4), the GP images revealed that the structure of gut epithelial cells was retained after Laurdan staining without any alterations on their cell morphology. It was possible to observe some changes in GP colors during the five development stages. For example, the GP coloring of the apical membrane shifted from

(middle degree of order) orange-green at 3 dpf to 6 dpf red (high order). In contrast, orange-green (middle degree of order) and green-blue (lowest order) colors intensity, 92

was observed in basolateral and intracellular membranes from 3 dpf till 6 dpf. Then the intensity of GP colors in apical, basolateral and intracellular membranes decreased gradually after 6 dpf till 11 dpf.

To obtain GP values, 10 larvae were used for each developmental stage, and an overall of ROI = 50 image regions were analyzed for each membrane at each development stage. The data in (Figure 3-5) showed that GP values of apical membrane of polarized epithelial cells in the gut from 3 to 11 dpf were significantly higher than of basolateral and intracellular membranes (p < 0.0001). Moreover, the GP values of basolateral membrane were significantly higher than intracellular membrane’s GP values (p < 0.0001) (Figure 3-4 and 3-5).

When compared over the course of development (Figure 3-8), the data revealed that membrane order of apical and basolateral membranes of polarized epithelial cells of the gut increased during zebrafish development stages from 3 dpf to 6 dpf while GP values decreased from 6 dpf to 8 dpf and 11 dpf. We observed that GP values of basolateral membrane at 6 dpf showed no significant differences with GP values at 8 dpf (p =

0.3319). Further, GP values of both apical and basolateral membranes at 3 dpf were significantly lower than GP values at 11 dpf (p < 0.0001) (Figure 3-8A). In summary, the GP analysis revealed that the membrane order of apical and basolateral membranes peaked at 6 dpf, and that at all stages of development, the apical membrane was significantly more ordered than the basolateral membrane, which was significantly more ordered than intracellular membranes.

3.3.2 Membrane order of epithelial cells in the kidney during development

Resembling a mammal’s kidney, the zebrafish kidney has two nephrons with a glomerulus, proximal tubules, distal tubules, and collecting ducts (Reimschuessel and

Ferguson 2006). The cell types of the pronephric glomerulus are also similar to that of higher vertebrate kidneys, such as podocytes, and polarized tubular epithelial cells, which display a projecting brush border (Drummond et al. 1998). These tubular epithelial cells are called pronephric tubules and formed a single layer of cube-like cells

(simple cuboidal epithelia) with relatively large nuclei. As in the gut, the apical membranes have a brush border face the lumen and the basolateral membranes correspond to the lateral borders (Figure 3-6). Kidney Pronephric ducts

IM Lumen BM

AM Pronephric tubules

Figure 3-6: GP image of zebrafish kidney. Image of kidney pronephron of zebrafish showing the pronephric tubules, which is simple cuboidal epithelia with a relative large nucleus. Apical membrane (AM) (highlighted in white), basolateral membrane (BM) (indicated by the yellow selection) and intracellular membrane (IM) (highlighted in blue).

The GP images of the kidney epithelial cells in Figure 3-6 indicate that the structure and morphology of the epithelial cells was not affected by Laurdan staining during all development stages. An increase in GP value in apical (mainly red GP coloring), basolateral (mainly orange-green GP coloring) and intracellular membranes (mainly green-blue GP coloring) was observed from 3 dpf till 6 dpf, which decreased gradually

after 6 dpf. To measure membrane GP, 10 larvae were used for each developmental stage, and a total of ROI = 50 image regions were analyzed for each membrane at each development stage.

Similar to what was found in the gut, at the five different development stages of the kidney, the GP values of the apical membrane of polarized epithelial cells were significantly higher than GP values of basolateral and intracellular membranes (p <

0.0001), and GP values of basolateral membrane were significantly higher than GP values of intracellular membrane (p < 0.0001) (Figure 3-5 and 3-8).

By comparing GP values of apical and basolateral membranes of polarized epithelial cells of the kidney during developmental stages, an increase in GP values of apical and basolateral membranes was noted during zebrafish development until GP values reached their peak at 6 dpf. Like in the gut, GP values of apical and basolateral membranes decreased gradually at 8 dpf and 11 dpf stages. When comparing GP values of apical and basolateral membranes at 3 dpf and 11 dpf, the data did not show any differences between these two developmental stages (p = 0.1473 and p = 0.0841, respectively) (Figure 3-8B).

3.3.3 Membrane order of epithelial cells in the liver during development

Zebrafish liver epithelial cells, hepatocytes, do not have a particular structure or a polyhedral shape, and these cells are normally arranged in acinar form; they are not distinctly arranged in cords, and the typical portal triads are not observable, as is common in mammal livers (Van Der Ven et al. 2003). In contrast to epithelial cells of the gut and kidney, hepatocytes have multiple apical and basolateral membranes. The apical membranes, sometimes called canalicular membrane, face the bile canaliculi (and 95

could not be seen in GP images), which is in contact with the outer environment, into which bile is secreted. The basolateral membranes face the sinusoids and also called the sinusoidal membrane (Decaens et al. 2008) while intracellular membranes are within the cells (Figure 3-7). Liver

Hepatocytes

IM AM BM

Sinusoids

Figure 3-7: GP image of zebrafish liver. Hepatocytes, the main epithelial cell type in the zebrafish liver, have no specific shape but have more than one apical and basolateral membrane. Apical membranes (AM) (highlighted in white) face the bile canaliculi, which cannot be seen in GP image, basolateral membranes (BM) (indicated by the yellow selection) face the sinusoids, and intracellular membrane (IM) (highlighted in blue).

The GP image in Figure 3-4 suggests that Laurdan staining did not induce any major changes in the liver epithelium since the cell morphology looked normal. As indicated by the GP colors, changes in membrane order of the apical (mainly red GP coloring), basolateral (with a predominate GP color of orange-green) and intracellular

(mainly green-blue GP coloring) occurred during the five development stages, which the highest membrane order observed at 6 dpf. For quantification of GP values, a total of ROI = 50 image regions for each developmental stage were selected from 10 larvae and analyzed for each membrane (Figure 3-4).

Similar to the data in gut and kidney described above, the GP values of apical membranes were significantly higher than GP values of basolateral and intracellular membranes (p < 0.0001). Also, GP values of basolateral membranes were significantly higher than those of intracellular membranes (p < 0.0001) (Figure 3-4 and 3-5).

Resembling kidney data, GP values of apical and basolateral membranes of polarized epithelial cells of the liver showed an increasing trend during zebrafish development until 6 dpf, then GP values gradually decreased from 6 dpf to 8 dpf and 11 dpf. GP values of apical membranes at 3 dpf and 11 dpf were not significantly different at these two development stages (p = 0.5343) while the GP values of the basolateral membrane at 3 dpf were significantly lower than at 11 dpf (p < 0.0001) (Figure 3-8C).

Figure 3-8: Changes in GP values of apical and basolateral membranes during zebrafish development of gut (A) kidney (B) and liver (C). In all three tissues and in both apical and basolateral membranes did the GP values peak at 6 dpf. n = 10 larvae were used for each developmental stage and a total of ROI = 50 image regions were analyzed for each membrane at each development stage. Symbols represent mean and SD. All data points were significantly different to each other (p < 0.05), except were indicate with ns (p > 0.05) (one-way ANOVA with Bonferroni post-test).

3.4 Effect of PFA fixation on GP values measurements

So far, all GP measurements were performed on live zebrafish larvae. It would be convenient to fix larvae for imaging and allow a direct comparison between GP measurements and immunostaining. To study the effect of fixation with 4% PFA on the

GP values measurements, I compared the GP values of apical, basolateral and intracellular membranes of polarized epithelial cells of gut, kidney, and liver in live and fixed larvae. Casper fish were mated and 6 dpf larvae labeled with Laurdan as described

in Section 2.2.1 and divided into two groups. In the first group, larvae were anesthetized and imaged live as described in Section 2.2.2. Larvae in the second group were fixed with 4% PFA overnight at 4°C followed by with PBS washing next day, and then imaged in the same manner as the live larvae; i.e. mounted in 1% agarose and under the identical microscopy settings. Membrane GP of epithelial cells of live and fixed larvae were quantified and compared.

Figure 3-9: Comparison of GP values of the apical (AM), basolateral (BM), and intracellular membranes (IM) between live and fixed larvae. GP values of apical (AM), basolateral (BM) and intracellular membranes (IM) of gut (A), kidney (B) and liver (C) of 6 dpf zebrafish larvae that were imaged live and after fixation with paraformaldehyde. Data are mean and standard deviations of 50 regions of interests obtained from 10 larvae. ns, not significant, p > 0.05; * p < 0.05; ** p < 0.01 between fixed and live larvae (Student t test). AM = apical membrane, BM = basolateral membrane and IM = intracellular membrane.

Figure 3-9 show that GP values of apical, basolateral membrane and intracellular membranes of gut, kidney, and liver of live zebrafish larvae at 6 dpf were either similar or higher than GP values of apical, basolateral and intracellular membranes of fixed larvae at 6 dpf. In the gut, GP values of apical of gut epithelial cells of live larvae were not significant different (p = 0.2811) from fixed larvae, while membrane GP of basolateral and intracellular membranes of fixed larvae were slightly lower than those of live larvae (p = 0.0149 and p = 0.0017, respectively) (Figure 3-9A). In the kidney, GP values of apical and basolateral membranes of kidney epithelial cells of fixed larvae were moderately lower than GP values of live larvae (p = 0.0014 and p = 0.0026,

respectively) while GP values of intracellular membrane of fixed larvae were significantly lower than GP values of live larvae (P< 0.0001) (Figure 3-9B). In the liver,

GP values of apical and basolateral membranes of liver epithelial cells of live and fixed larvae were similar (p = 0.7406, and p = 0.7729 respectively), while GP values of intracellular membrane of fixed larvae were significantly lower than membrane GP of live larvae (p < 0.0001) (Figure 3-9C). Fixation with 4% PFA seemed to be slightly affected the measurements of GP values between live and fixed 6 dpf larvae causing a decrease in membrane order in all three examined membranes (Figure 3-9). However, the differences between apical, basolateral and intracellular membranes in all three tissues were maintained with apical membranes remaining significantly more ordered than basolateral and intracellular membranes and GP values of basolateral membrane significantly higher than GP values of intracellular membrane (Figure 3-10).

Figure 3-10: GP values of apical (AM), basolateral (BM), and intracellular membranes (IM) at fixed 6 dpf larvae. GP values of apical, basolateral and intracellular membranes of gut (A), kidney (B) and liver (C) of fixed larvae. Data are mean and standard deviations of 50 regions of interests obtained from 10 larvae. **** indicates statistical difference of p < 0.0001 (one-way ANOVA with Bonferroni post-test). AM = apical membrane, BM = basolateral membrane, and IM = intracellular membrane.

3.5 Conclusion and Discussion

The zebrafish strain Casper presented itself as the appropriate choice to investigate membrane order during different development stages due to their transparency, which remain until adulthood. We developed and demonstrated that Laurdan microscopy on

Casper zebrafish larvae allows us to image and quantify membrane order in living and fixed organisms. The work presented here may facilitate further membrane order studies in other zebrafish tissues.

The aim of this chapter was to investigate membrane order of polarized epithelial cells of the gut, kidney, and liver during different development stages in zebrafish. We verified that the apical membrane of polarized epithelial cells was significantly more ordered than the basolateral membranes in gut, kidney and liver at all five developmental stages (Figure 3-7), which is consistent with what was found in studies with cultured epithelial cells (Vieira et al. 2006) and in whole living organism (Owen et al. 2010).

We noticed that there was an increase and then decrease in membrane order of epithelial cells of gut, kidney, and liver during the five development stages. To quantify these changes, we measured the GP value in regions corresponding to the apical and basolateral membranes. After comparing GP values of apical and basolateral membranes in gut, kidney, and liver, across the five development stages, the increase in membrane order occurred from 3 dpf to 6 dpf with particularly high membrane orders recorded at 6 dpf for the three tissues. After 6 dpf the membrane order decreased through 8 dpf till it reached the lowest at 11 dpf (Figure 3-8).

100

In summary, membrane order of these cells was significantly remodeled during development and peaked at 6 dpf in both apical and basolateral membranes. The possible explanation for this is that zebrafish larvae commence feeding and tissues are almost completely developed, especially the digestive tract, at about 5 dpf (Flynn et al.

2009). Therefore, it may that the increasing of membrane order of epithelial cells from 3 dpf to 6 dpf is related to developing of the tissues. Since membrane order reaches its peak at the stage when larvae start feeding and tissues are nearly developed completely, the decreasing membrane order of polarized epithelial cells at 11 dpf may reflect the steady state condition of adult fish. It is possible that the remodeling of membrane order during development plays a functional role and aids polarized epithelial cells in functions such as in sorting and trafficking of lipids and proteins via the secretory and endocytic pathways (Bretscher and Munro 1993; Simons and Ikonen 1997).

Here, multi-photon microscopy with Laurdan proved to be a useful technique for studying membrane order of epithelial cells in vivo. We revealed that Laurdan could by itself mark zebrafish tissues as individual cells were clearly visible, and apical, basolateral and intracellular membranes could easily be recognized (Figure 3-4) without any transient expression of mCherry in zebrafish embryos, which was used before to label the region of interest in a previous Laurdan study (Owen et al. 2010). We verified that Laurdan did not affect zebrafish development as the 3 dpf treated larvae, with two different Laurdan concentrations, developed normally without affecting their viability or behavior. In addition, our data showed that fixing zebrafish larvae with 4% PFA affected the absolute GP values but maintained the difference in membrane order within cellular membranes. Hence the data indicate that it possible to proceed in our further

101

investigations in fixed larvae and ask whether sterol manipulations of intact zebrafish embryos affect membrane order and localization of polarity proteins.

102

Chapter 4: Effect of sterol manipulations on membrane order and polarity protein localization in polarized epithelial cells in vivo

4.1 Introduction

In order to understand the functional roles of the membrane order in model membranes and live cells, it is common to manipulate the sterol content and investigate whether altered sterol levels affect the function of interest. Due to the important role of cholesterol in lipid raft formation and stabilization, cholesterol levels are often increased or decreased with methyl-β-cyclodextrin (mβCD). Cholesterol is the key element that maintains cell membranes and adjusts membrane fluidity over the range of physiological temperatures, as it increases lipid packing via interaction with the phospholipid fatty-acid chains (Bloch 1991; Sadava et al. 2011). For example, several studies in model membranes revealed the influence of cholesterol on the stability and organization of (lo) and (ld) phases (Silvius 2003; Crane and Tamm 2004). It has also been proposed that cholesterol has pivotal implication in the establishment of liquid- ordered and liquid-disordered phase in the plasma membrane of mammalian cells, which supposedly act as platforms for receptors proteins and second messenger molecules (Simons and Ikonen, 1997; Incardona and Eaton 2000).

Since cholesterol has an ability to modify the physicochemical features of model and cellular membranes, sterol manipulations may also affect membrane protein localization and function. It has been found that a number of integral membrane proteins such as ion channels, membrane receptors, and enzymes are functionally affected by

103

physical differences in the surrounding lipid bilayer. Additionally, cholesterol is a key factor for the function of raft-associated proteins (Incardona and Eaton 2000).

It has been found previously that some proteins are dependent on direct binding to cholesterol such as VIP21/caveolin-1 which when binds to cholesterol to establish caveolae and function in membrane trafficking (Murata et al. 1995). In erythrocyte plasma membranes, which contain a set of distinct proteins and high cholesterol levels, a depletion of cholesterol showed a disruption of raft-associated protein localization in the plasma membrane that confirmed the fundamental role of cholesterol in proteins assembly into rafts domains. On the other hand, increased cholesterol levels induced a defect in targeting of GPI-anchored proteins to rafts, but this increase in cholesterol levels did not increase raft association of proteins, proposing that GPI-anchored proteins do not bind directly to cholesterol (Samuel et al. 2001).

Polarity proteins are essential membrane proteins in epithelial cells and are responsible for establishing and maintaining cell polarity. Atypical protein kinase C

(aPKC) is one of these important proteins that from a network with other polarity proteins, namely Par3 and Par6. aPKC in Drosophila is localized at the apical region of embryonic epithelial cells and neuroblasts (Petronczki and Knoblich 2001) while in mammals, it is localized asymmetrically at TJ (tight junctions) of epithelial cells together with the protein ZO-1 (Izumi et al. 1998; Suzuki et al. 2001). In zebrafish, aPKCs was found at the apical domain of polarized epithelial cells of the swim bladder, esophagus and intestine in addition to a subapical localization together with ZO-1 protein in retinal neuroepithelium at 32 hpf (Horne-Badovinac et al. 2001). A correlation between membrane order and polarity proteins has not yet been investigated.

104

In this chapter, I will examine the effect of different sterol treatments such as cholesterol extraction from the cellular membrane and sterol enrichment, on membrane order of epithelial cells and the apical localization of aPKC. The data may clarify whether there is a correlation between membrane order and polarity proteins.

To achieve this aim, 7-ketocholesterol (7KC) and methyl-β-cyclodextrin (mβCD) were employed in this chapter. The oxysterol 7-ketocholesterol (7KC) is an oxygenated derivative of cholesterol, and generally has the same cellular processing and intracellular trafficking of cholesterol. 7KC differs from cholesterol by an additional ketone group, which stand out perpendicular from the planar cholesterol ring. 7KC is known to prevent the formation of liquid-ordered domains in model membranes by hindering the compaction of saturated acyl chains in phospholipid bilayers (Massey and

Pownall 2005). Rentero et al. previously used 7KC as an efficient compound to disrupt membrane order in cultured T cells. At low concentrations, 7KC had no significant impact on cell viability and proximal signaling activities but severely impaired T cell activation by preventing the formation of signaling complexes at the cell membrane

(Rentero et al. 2008). In a related study, Owen et al. revealed that 7KC decreased the membrane order in the immunological synapse (IS) and to a lesser extent, of intracellular membranes in T cells. Even after 90 min treatment with 7KC, T cells retained their normal morphology and the decreased membrane order did not influence the capability of T cells to adhere to activating surfaces (Owen et al. 2010). To investigate the relationship between fenestrations transcellular holes, which facilitate substrates movement between blood and the extravascular compartment, in endothelial cells and membrane rafts in liver sinusoidal endothelial cells, 7KC was employed and

105

shown to increase lipid-disordered membrane in these cells. 7KC treatment increased the number and diameter of fenestrations in vitro while in vivo, 7KC increased only the diameter of fenestrations (Quinn et al. 2012). In addition, 7KC was used to present a direct proof that the plasma membrane of intact, live cells are comprised of a sub- resolution combination of ordered and disordered lipid domains. While in non-treated cells, ordered membrane covered ~76% of the plasma membrane, treating HeLa cells with 25 μM 7KC for 30 min reduced the membrane ordered coverage to ~25% in these cells. 7KC treatments did not reduced the fluidity of disordered regions or created a third lipid phase (Owen, et al. 2012).

Cyclodextrin are cyclic oligosaccharides made of α-(1–4)-linked D-glycopyranose units, which are a primary product of starch degradation. Methyl-β-cyclodextrin

(mβCD) is one of the most effective compounds among cyclodextrin to extract cholesterol from the cell membrane (Davis and Brewster 2004). This compound has been known as a highly effective transporter for hydrophobic drugs because it contains a hydrophobic cavity, which can encapsulate different hydrophobic molecules (Pitha et al. 1988). MβCD have the highest affinity for cholesterol among other cyclodextrins, and is considered the most effective way to extract cholesterol from membranes (Ohtani et al. 1989; Ohvo and Slotte 1996). MβCD is also soluble in water and form soluble inclusion complexes with cholesterol, thus increasing the solubility of cholesterol in aqueous solution. In summary, mβCD can be used to shuttle cholesterol between cells and solution; when added as cholesterol-free mβCD it generally removes cholesterol from cells, when added as mβCD/cholesterol complex, it can increase the cellular cholesterol content (Pitha et al. 1988).

106

The role of membrane order in cellular processes such as T cell activation can be investigated by using cyclodextrin, particularly mβCD, to deplete cholesterol from the plasma membrane of cultured cells and disrupt membrane order of these cells

(Kabouridis et al. 2000; Rentero et al. 2008). On the other hand, in order to study the role of rafts in apical transport in epithelial Madin-Darby canine kidney (MDCK) cells, cholesterol was extracted from the cell membranes by mβCD and the secretion of three different classes of proteins from the apical and basolateral membranes determined. The outcome of this study emphasized that cholesterol depletion modified the apical to basolateral secretion ratio by decreasing apical transport without affecting sorting processes in MDCK cells (Prydz and Simons 2001). Unfortunately, it has been found that at high concentrations, mβCD may cause adverse effects such as loss of cell viability, morphology changes, loss of membrane fatty acids, modifications in proteins, barrier dysfunction and cytoskeleton reorganization (Hinzey et al. 2012).

In this chapter, I asked whether membrane order can be manipulated by enrichment and deplete cholesterol in polarized epithelial cells in intact zebrafish and whether the sterol manipulations will affect the apical targeting of polarity proteins. The first section of this chapter describes the sterol manipulations with 7KC, cholesterol-mβCD complexes, and mβCD to investigate their effects on membrane order of epithelial cells in the gut, kidney, and liver and investigate changes on the cellular structures after treatments. In the second part, I investigated whether the membrane order recovers after sterol manipulations by keeping the treated larvae alive for two additional days, followed by the quantification of the GP values of the apical, basolateral and intracellular membranes of epithelial cells of the three tissues.

107

In the last part, I immunostained larvae after sterol manipulations to study the effect of sterol treatments on the apical localization of the polarity protein aPKC and whether its apical localization is recovered after two days post sterol manipulations.

4.2 Sterol manipulations of intact larvae

To examine whether membrane order affects the distribution of polarity proteins in polarized epithelial cells, I first aimed to manipulate the membrane order with different sterol treatments without disturbing the overall tissue architecture. The preparation of sterol-mβCD complexes is described in the Materials and Methods chapter, Section 2.4.

4.2.1 Conditions of sterol manipulations of intact larvae

To manipulate the membrane order of epithelial cells of gut, kidney and liver, three groups of 6 dpf larvae were treated as follow: 1) larvae were treated with 100 µM 7KC- mβCD, by adding 10.06 µl of 4.97 mM 7KC-mβCD to 1 ml E3 medium, for 30 min at

28°C with the goal to increase the fluidity of cellular membranes; 2) larvae were treated with 100 µM cholesterol- mβCD, by adding 17.7 µl of 5.64 mM cholesterol-mβCD to 1 ml E3 medium, for 30 min at 28°C to increase cholesterol levels; 3) larvae were treated with 2.5 mM mβCD, by adding 0.0033 g of mβCD to 1 ml E3 medium, for 40 min at

28°C to extract cholesterol. As a fourth and Control group, larvae were not treated with any compound and kept in E3 medium at 28°C.

These concentrations were chosen after performing a series of experiments to figure out the appropriate concentration of 7KC- mβCD, cholesterol-mβCD, and mβCD complexes that disrupt membrane order of the epithelial cells without causing larvae death. Using larvae at 6 dpf, it was found that at higher concentrations and longer 108

incubation times as those stated above, larvae disintegrated and individual cells were no longer visible after Laurdan staining.

4.2.2 Effect of sterol manipulations on tissue morphology

We asked whether epithelial cells retained their architecture and the tissue morphology after enrichment with 7KC and manipulations of the cholesterol level and whether the sterol treatment conditions were suitable for membrane order measurement.

The sterol levels of 6 dpf larvae were manipulated as described above, fixed, stained for

ZO-1, WGA, CTxB and DAPI and imaged by confocal microscopy as described in

Section 2.3.1 and 2.3.2.

109

Figure 4-1: Tissue morphology of gut (A), kidney, (B) and liver (C) in sterol treated 6 dpf zebrafish larvae. 6 dpf larvae was left untreated (Control), treated with 7-ketocholesterol complexed to mβCD (7KC, 100 μM, 30 min) or mβCD/cholesterol (cholesterol, 100 µM, 30 min) or depleted of cholesterol with methyl-ß-cyclodextrin (mβCD, 2.5 mM, 40 min), fixed and labelled with antibodies against the tight junction marker ZO-1 (red), WGA (yellow) to stain the cell surface, CTxB (green) to label GM1-rich membranes and DAPI (blue) to visualize cell nuclei. Scale bar in merged imaged = 20 µm; scale bar in zoomed merged images = 5 µm. 110

Figure 4-1 displays the confocal images of ZO-1, WGA, CTxB, and DAPI in the gut, kidney, and liver after 7KC, cholesterol, and mβCD treatments in comparison to

Control larvae. In gut epithelial cells treated with 7KC, cholesterol and mβCD (Figure

4-1A), the normal cellular structure and tissue organization was retained. ZO-1 (shown in red) had a punctate staining that was localized to cell-cell interfaces (nuclei were stained with DAPI and are shown in blue). In the merged images, it is apparent that ZO-

1, as expected, stained the tight junction of epithelial cells. Since the ZO-1 staining pattern was similar in all conditions, it was concluded that the formation of tight junctions of gut was not affected by the sterol manipulations. WGA (shown in yellow) stains the gut epithelial cells surface. In the merged images, the WGA showed an even staining under all conditions without accumulating in specific regions suggesting that the structure of gut epithelial cells was not affected by the sterol manipulations. CTxB

(shown in green) labels GM-1 rich membranes, which are mainly localized in apical regions. In the merged images, it is visible that CTxB stained the apical membrane of epithelial cells even in sterol-manipulated larvae. In conclusion, the sterol manipulations did not appear to affect the tissue architecture of the gut.

Figure 4-1B shows confocal images of the kidney that epithelial cells indicating that they maintained their normal morphological structure and tissue organization after treatment with 7KC, cholesterol-mβCD, and mβCD. As in the gut, ZO-1 (shown in red) staining was restricted to the cell-cell interfaces and away from the nuclei stained with

DAPI (shown in blue) suggesting that tight junctions were maintained in all conditions.

Similarly, WGA (shown in yellow) stained the kidney epithelial cells surface in a homogenous fashion and the cell arrangement in sterol-treated larvae looked similar to

111

Control tissue. CTxB (shown in green) marked the GM-1 rich membranes, which mainly localized at the brush border of kidney epithelial cells. In the merged images, it can be seen that CTxB stained the apical membranes of epithelial cells. Since the ZO-1,

WGA, and CTxB staining were similar in all conditions, it was concluded that the kidney epithelial cells were not distorted by the sterol manipulations.

Figure 4-1C shows that the architecture of liver epithelial cells was morphologically maintained after sterol manipulations. The nuclei stained with DAPI

(shown in blue) and tight junction stained with ZO-1 (shown in red) appeared normal.

WGA (shown in yellow) stained the entire surface of liver epithelial cells surface and

CTxB (shown in green) marked GM-1-rich membranes of apical membranes of liver epithelial cells. Hence, the staining of ZO-1, WGA, and CTxB under all three sterol manipulations was comparable to Control liver emphasizing that sterol manipulations did not affect the cellular organization of the liver.

4.2.3 Membrane order in sterol-manipulated larvae

To assess whether 7KC, cholesterol, and mβCD treatments changed the membrane order of polarized epithelial cells in the gut, kidney and liver of 6 dpf zebrafish larvae, a batch of larvae from each of the four groups (Control, 7KC, cholesterol-mβCD, and mβCD) were stained with Laurdan as in Section 2.2.1, fixed with 4% PFA and imaged as in Section 2.2.2. GP values of the plasma membrane GP in the gut, kidney, and liver were quantified and presented in Figure 4-2. For each group, 15 images were acquired from 15 larvae for each tissue, and from each tissue five regions of interest (ROI) were selected for the apical, basolateral membrane and intracellular membranes in each larva.

112

** ** ** **

Figure 4-2: Membrane order in sterol treated 6 dpf zebrafish larvae. Larvae at 6 dpf were left untreated (Control), treated with 7-ketocholesterol (7KC 100 µM for 30 min), cholesterol-mβCD (cholesterol 100 µM for 30 min) or methyl-β-cyclodextrin (mβCD 2.5 mM for 40 min), stained with Laurdan and GP images of gut (A), kidney (B) and liver (C) obtained. GP values were quantified for epithelial cells in the gut (D), kidney (E) and liver (F). In A-C, scale bar = 20 µm. In D-F, 15 larvae were used with a total of ROI = 75 were analyzed for each membrane and each treatment. Data are mean and standard deviations. **** indicates a statistical difference of p < 0.0001 (one-way ANOVA with Bonferroni post-test). AM = apical membrane, BM = basolateral membrane, and IM = intracellular membrane. ** indicate for gut lumen. 4.2.4 Membrane order of epithelial cells in the gut

In the GP images of gut epithelial cells (Figure 4-2A), cells were clearly visible in

Control and all sterol manipulation GP images. Cells in the gut appeared structurally normal, and neither the sterol manipulations nor Laurdan staining affected the cell morphology. The GP images indicate sterol manipulations with 7KC, cholesterol and mβCD decreased membrane order in the gut compared to Control tissue since the coloring was shifted from red (representing high membrane order) to predominately green (representing low membrane order).

113

The shift towards more fluid membranes was confirmed by comparing the GP values of treated cells versus Control. The data revealed that the GP values decreased significantly in apical, basolateral and intracellular membranes in larvae treated with

7KC, mβCD and cholesterol compared to Control (p < 0.0001) (Figure 4-2D). It was surprising that treatment with cholesterol-mβCD reduced membrane order to a similar extent than 7KC and mβCD treatment. It was expected that cholesterol-mβCD would increase cholesterol level and possibly membrane order. It is possible that cholesterol- mβCD resulted in cholesterol re-distribution and/or net cholesterol reduction, which could decrease membrane order in polarized epithelial cells.

All sterol treatments significantly decreased the membrane order of apical, basolateral and intracellular membranes of the gut epithelial cells to almost the same level (Figure 4-2D). The exception was treatment with cholesterol-mβCD, which reduced the GP values of basolateral membrane more than mβCD treatment (p =

0.0393).

As described in Chapter 3, Section 3.3 (Figure 3-5), apical membranes were significantly more ordered in gut, kidney, and liver epithelial cells than basolateral membranes, which in turn were significantly more ordered than intracellular membranes.

The difference in order between membranes was maintained in the gut after 7KC (p <

0.0001) apical versus basolateral membrane, (p < 0.0001) basolateral versus intracellular membranes, cholesterol-mβCD (p < 0.0001) apical versus basolateral membrane, (p < 0.0001) basolateral versus intracellular membranes) and mβCD (p <

0.0001) apical versus basolateral membrane, (p < 0.0001) basolateral versus intracellular membranes treatment (Figure 4-2D).

114

4.2.5 Membrane order of epithelial cells in the kidney

GP images of epithelial cells in the kidney at 6 dpf (Figure 4-2B) show that cellular structure was maintained in all sterol-treated larvae compared to Control. Like in the gut, it was noticed that membrane order in epithelial cells treated with 7KC, cholesterol- mβCD, and mβCD was reduced relative to Control larvae.

GP values of the kidney epithelial cells of treated larvae were significantly decreased in apical, basolateral and intracellular membranes compared to Control (p <

0.0001) (Figure 4-2E). Surprisingly, the data demonstrated that the cholesterol-mβCD treatment had the highest impact on GP values. The GP values of apical membrane of epithelial cells treated with cholesterol were slightly lower than GP values of epithelial cells treated with 7KC (p = 0.0298). In addition, GP values of basolateral membrane of epithelial cells treated with cholesterol-mβCD were slightly lower than those treated with 7KC and mβCD (p = 0.0206 and p = 0.0225 respectively). Finally, GP values of intracellular membrane of epithelial cells treated with cholesterol-mβCD were significantly lower than GP values of intracellular membrane of epithelial cells treated with 7KC and mβCD (p < 0.0001) (Figure 4-2E).

Despite the reduction in membrane order, sterol-manipulated kidney epithelial cells maintained differences between apical, basolateral and intracellular membranes in terms of membrane order. In all sterol-treated larvae, apical membrane remained more ordered than basolateral membrane (p < 0.0001) and basolateral membranes remained more ordered than intracellular membrane (p < 0.0001) (Figure 4-2E).

115

4.2.6 Membrane order of epithelia cells in the liver

Similar to the observation in the gut and kidney, the GP images of the epithelial cells of the liver (Figure 4-2C) indicate that sterol manipulations and Laurdan staining did not disrupt the overall cellular organization of liver epithelial cells. Furthermore, the

GP images indicate a decreasing in membrane order of apical, basolateral and intracellular compared to Control larvae. The GP values of apical, basolateral and intracellular membranes of polarized epithelial cells of zebrafish liver were significantly decreased compared to Control (p < 0.0001) (Figure 4-2F). Here, we noticed that 7KC has the highest impact on decreasing membrane order of apical and basolateral membranes. The data revealed that GP values of apical membrane of epithelial cells treated with 7KC were slightly lower than GP values of epithelial cells treated with cholesterol-mβCD and mβCD (p = 0.0257 and p = 0.0364 respectively) (Figure 4-3C).

In basolateral membrane, the GP values of epithelial cells treated with 7KC were moderately lower than GP values of cells treated with cholesterol-mβCD and mβCD (p

= 0.0004 and p = 0.0021 respectively) (Figure 4-2F). It appears that all three treatments had a similar effect on decreasing GP values of intracellular membrane (Figure 4-2F).

Again, the differences in membrane order between apical, basolateral and intracellular membranes remained after sterol treatment in liver epithelial cells.

Membrane order of apical membrane still significantly higher than basolateral membrane, and membrane order of basolateral membrane also remained higher than intracellular membrane in all sterol-treated larvae (Figure 4-2F).

116

In order to discover which tissue was most affected by the sterol treatments at 6 dpf stage, additional comparisons were done between the GP values of three membranes of gut, kidney and liver.

Apical membrane

Within 7KC treated larvae, the apical membranes of liver were the most affected

after sterol manipulations; GP values of apical membranes of liver epithelial cells

were significantly lower than GP values of apical membrane of kidney epithelial

cells (p < 0.0001), and slightly lower than those of gut epithelial cells (p = 0.0026).

Within cholesterol treated larvae, GP values of liver and kidney epithelial cells

were slightly lower than GP values of gut epithelial cells (p = 0.0014 and p =

0.0041 respectively), and GP values of epithelial cells of the liver showed no

differences with those of kidney (p = 0.6557). The effect of mβCD was the same on

the three tissues as GP values showed no significant differences between all of the

three tissues (Figure 4-2D, E & F). In summary, at 6 dpf, apical membranes of liver

epithelial cells were the most affected with 7KC and cholesterol-mβCD treatments

while the three tissues were affected at the same level with mβCD complex.

Basolateral membrane

In larvae treated with 7KC, the basolateral membranes in gut and liver epithelial cells were more affected than basolateral membranes in kidney epithelial cells. The data revealed that GP values of basolateral membrane of gut and liver epithelial cells were slightly lower than GP values of kidney epithelial cells (p = 0.0004 and p = 0.0068 respectively) (Figure 4-2D, E & F). Within larvae treated with cholesterol-mβCD, GP values of basolateral membranes of gut epithelial cells were significantly lower than GP 117

values of liver epithelial cells (p < 0.0001), and slightly lower than GP values of basolateral membranes of kidney epithelial cells (p = 0.0351) (Figure 4-2D, E & F).

Within mβCD treated larvae, GP values of basolateral membrane of gut epithelial cells were significantly lower than those of kidney and liver epithelial cells (p = 0.0030 and p

= 0.0007, respectively) (Figure 4-2D, E & F). In summary, basolateral membranes of gut epithelial cells were the most affected by the three sterol modifications.

Intracellular membrane

Within larvae treated with 7KC, GP values of intracellular membranes of gut and liver epithelial cells significantly decreased compared to GP values of those of kidney epithelial cells (p < 0.0001) (Figure 4-2D, E & F). In larvae treated with cholesterol- mβCD, the comparison showed no significant differences between GP values of intracellular membrane of epithelial cells of gut, kidney, and liver.

In mβCD-treated larvae, GP values of intracellular membrane of epithelial cells of liver and gut were significantly lower than those of kidney (p < 0.0001 and p = 0.0006 respectively) (Figure 4-2D, E & F). In summary, gut and liver epithelial cells were the most affected by 7KC and mβCD treatments while all three tissues were equally affected with cholesterol-mβCD complexes.

4.3 Recovery of membrane order after sterol manipulations

After examining the impact of sterol manipulations on the membrane order of polarized epithelial cells of 6 dpf larvae, I proceeded to find out whether the membrane order of these epithelial cells was retained after further development. Therefore, two groups of 4 dpf larvae were treated with 7KC, cholesterol-mβCD, or mβCD as in

118

Section 4.2.1. A batch from each sterol treatment, including Control larvae, were fixed immediately and their membrane order measured after Laurdan staining as in Section

2.2.1 and Section 2.2.2. The other batches of treated and Control larvae were allowed to grow in E3 medium at 28°C till 6 dpf and then stained with Laurdan, fixed and imaged.

For details of the workflow are shown in Figure 4-3.

Figure 4-3: Illustration of sterol manipulation experiment. Two groups of 4 dpf larvae were treated with 7KC, cholesterol, and mβCD or left untreated and membrane order and aPKC apical localization were examined for one group at that developmental stage. One group of each of the treated larvae was allowed to develop until 6 dpf and membrane order and aPKC apical localization examined at 6 pdf.

At 4 dpf stage, from each condition (Control, 7KC, cholesterol-mβCD, and mβCD),

10 images were acquired from 10 larvae for each tissue, and from each larvae, 5 ROI were selected randomly for apical, basolateral and intracellular membranes for each gut, kidney and liver as described in Section 2.2 in Materials and Methods chapter.

119

** ** **

** **

Figure 4-4: Membrane order of sterol-manipulated larvae at 4 pdf and after a recovery period of two additional days (6 pdf). Larvae at 4 dpf were left untreated (Control), treated with 7-ketocholesterol (7KC 100 µ for 30 min), enriched in cholesterol (cholesterol 100 µ for 30 min) or depleted cholesterol with methyl-β-cyclodextrin (mβCD 2.5 mM for 40 min). A second group of larvae was tested at 4 dpf and allowed to develop to 6 dpf and then imaged. (A-B) GP images of gut, kidney, and liver of 4 dpf (A) and 6 dpf larvae (B) are shown. (C). GP values quantified for epithelial cells in the gut, kidney and liver at 4 dpf (C) and 6 dpf (D). In A and B, scale bar = 20 µm. In C and D, 10 larvae were used with a total of ROI = 50 were analyzed for each membrane and for each condition. Data are mean and standard deviations. ns, not significant, p > 0.05; * p < 0.01; *** p < 0.001; **** p < 0.0001 relative to Control (one-way ANOVA with Bonferroni post-test). In D, indicates a statistical difference of p < 0.0001 between GP values of 4 dpf and 6 dpf larvae (Student t-test). AM = apical membrane, BM = basolateral membrane, and IM = intracellular membrane. ** indicate for gut lumen. 120

4.3.1 Membrane order of epithelial cells in the gut, kidney and liver at 4 dpf

As was observed in 6 dpf larvae with sterol manipulations (Figure 4-2A), the cellular architecture and tissue configuration of gut, kidney and liver were also clearly not influenced by the sterol treatments at 4 dpf (Figure 4-4A). Obvious changes in GP colors (red, orange-green and green–blue) of the apical, basolateral and intracellular membranes compared to untreated larvae suggested that the membrane order was reduced in all three membranes in gut, kidney and liver after sterol manipulations

(Figure 4-4A).

Treating 4 dpf larvae with 7KC, cholesterol-mβCD and mβCD also significantly reduced the GP values of apical, basolateral and intracellular membranes of polarized epithelial cells of the gut, kidney, and liver comparing to Control (p < 0.0001). The GP values of apical membrane remained significantly higher than basolateral and intracellular membranes (Figure 4-4C).

Although all sterol treatments significantly decreased the membrane order of apical, basolateral and intracellular membranes in gut epithelial cells, further comparisons revealed that mβCD decreased GP values of apical, basolateral and intracellular membranes more than 7KC and cholesterol-mβCD complexes. It was found that GP values of apical membrane of gut epithelial cells treated with mβCD were slightly lower than GP values of those treated with 7KC (p = 0.0071), and significantly lower than those treated with cholesterol-mβCD (p < 0.0001) (Figure 4-4C). GP values of basolateral membrane of epithelial cells treated with mβCD were slightly lower than GP values of those treated with 7KC and cholesterol-mβCD (p = 0.0002 and p = 0.0245 121

respectively) (Figure 4-4C). Further, GP values of intracellular membrane of gut epithelial cells treated with mβCD were significantly lower than GP values of epithelial cells treated with 7KC (p < 0.0001) and lower than GP values of epithelial cells treated with cholesterol-mβCD (p = 0.0004) (Figure 4-4C).

Like in the gut, mβCD reduced the membrane order of apical, basolateral and intracellular membranes of kidney epithelial cells more than that of 7KC and cholesterol-mβCD complexes. Further, GP values of apical membranes of kidney epithelial cells treated with mβCD were slightly lower than GP values of those treated with 7KC (p = 0.0049), and significantly lower than epithelial cells treated with cholesterol-mβCD (p < 0.0001). In addition, GP values of basolateral membrane of epithelial cells treated with mβCD were slightly lower than GP values of those treated with 7KC (p = 0.0002), and significantly lower than GP values of epithelial cells treated with cholesterol-mβCD (p < 0.0001). In a similar manner, GP values of intracellular membrane of epithelial cells treated with mβCD were slightly less than GP values of epithelial cells treated with 7KC (p = 0.0099) and significantly lower than GP values of epithelial cells treated with cholesterol (P<0.0001) (Figure 4-4C).

In summary, mβCD had the highest effect on decreasing GP values of the three membranes in the kidney.

In liver epithelial cells, the data illustrated that mβCD and 7KC complexes decreased the membrane order of apical, basolateral membranes to a similar level, and all three sterol treatments equally reduced membrane order of intracellular membrane.

GP values of apical membrane of liver epithelial cells treated with mβCD and GP values of those treated with 7KC (p = 0.9764) were both significantly lower than epithelial

122

cells treated with cholesterol-mβCD (p < 0.0001) (Figure 4-4C). No differences were found between GP values of basolateral membrane of epithelial cells treated with mβCD and those treated with 7KC (p = 0.9094). GP values of basolateral membrane of epithelial cells treated with mβCD were significantly lower than GP values of epithelial cells treated with cholesterol-mβCD (p < 0.0001) (Figure 4-4C). No differences were observed between GP values of intracellular membrane of epithelial cells treated with mβCD and GP values of epithelial cells treated with 7KC and cholesterol-mβCD (p =

0.2416 and p = 0.1042, respectively) (Figure 4-4C).

4.3.2 Membrane order of gut, kidney and liver epithelial cells post recovery (6 dpf)

Two days after sterol treatment, at 6 dpf, GP images of the gut, kidney, and liver revealed that epithelial cells remained morphologically normal without any visible changes. The GP coloring indicates that apical, basolateral and intracellular membranes

GP values in treated larvae only slightly decrease comparing to Control (Figure 4-4B).

The GP colors of apical, basolateral and intracellular membranes at 6 dpf post recovery indicate an increase in membrane order compared to larvae at 4 dpf (Figure 4-4A).

Quantification of GP images at 6 dpf disclosed that GP values of apical, basolateral and intracellular membranes epithelial cells of gut, kidney and liver were still significantly reduced by the sterol manipulations, even after the recovery period (p < 0.0001)

(Figure 4-4D). However, a recovery of the membrane order in apical, basolateral and intracellular membranes could also be observed in gut epithelial cells. This was confirmed by comparing the GP values of apical, basolateral and intracellular membranes at 4 dpf with the GP values recorded at 6 dpf. This comparison revealed that

123

there was a significant increase in GP values of apical, basolateral and intracellular membranes from 4 dpf to 6 dpf (p < 0.0001) in the gut (Figure 4-4B), which suggested that there a membrane recovery occurred during the two days of development post sterol manipulations.

GP values were calculated for the kidney epithelial cells and compared to Control.

The GP values of apical membrane of larvae that was treated with 7KC at 4 dpf was no longer significantly different to Control post recovery (p = 0.1447). Apical membrane of epithelial cells treated with cholesterol-mβCD and mβCD had a slight decrease in GP value compared to Control after the recovery period (p = 0.0046 and p = 0.0004, respectively) (Figure 4-4D). GP values of basolateral membrane of kidney epithelial cells post-treated with 7KC and recovery showed a small decrease in GP value compared to Control (p = 0.0331), while a significant decrease in GP values was observed in basolateral membrane of kidney epithelial cells treated with cholesterol- mβCD and mβCD compared to Control post recovery (p < 0.0001) (Figure 4-4D). In intracellular membrane, GP values of kidney epithelial cells treated with 7KC and cholesterol-mβCD post-recovery showed a slight decrease compared to Control (p =

0.0016, p = 0.0214 respectively), and GP values of kidney epithelial cells treated with mβCD at 6 dpf displayed more of a decrease (p = 0.0002) than those treated with 7KC and cholesterol-mβCD at 6 dpf (Figure 4-4D). In summary, it was noticed that the membrane order of apical and basolateral membranes of kidney epithelial cells treated with 7KC, and membrane order of apical and intracellular membranes of kidney epithelial cells treated with cholesterol-mβCD had almost totally recovered. To a less degree, membrane order of apical and intracellular membranes of epithelial cells treated

124

with mβCD had also recovered (Figure 4-4D). In addition, comparing the GP values of apical, basolateral and intracellular membranes at 4 dpf with the GP values at 6 dpf confirmed the recovery of membrane order 6 dpf (p < 0.0001) (Figure 4-4D).

GP values of apical membrane of liver epithelial cells treated with 7KC and cholesterol-mβCD were slightly lower than in Control cells (p = 0.0008, p = 0.0005 respectively) while in liver epithelial cells treated with mβCD, significant differences in

GP values of apical membrane were observed comparing to Control (p < 0.0001)

(Figure 4-4D). In liver epithelial cells treated with cholesterol-mβCD, the GP values of basolateral membrane showed a slight decrease compared to Control (p = 0.0212) but

GP values of epithelial cells treated with 7KC and mβCD were significantly decreased

(p < 0.0001) (Figure 4-4D). In contrast, GP values of intracellular membrane of liver epithelial cells treated with cholesterol-mβCD showed no differences with the Control

(p = 0.0513), whereas a significant decrease were observed in liver epithelial cells treated with 7KC and mβCD (p < 0.0001) (Figure 4-4D).

125

Figure 4-5: Recovery of membrane order after sterol modification. Δ GP values obtained from the differences between GP values of apical (AM) and basolateral (BM) membranes in sterol-treated larvae at 4 dpf against their untreated control versus differences of GP values of sterol treated larvae at 6 dpf larvae against their untreated control. Data are mean and standard deviations from 50 regions in 10 larvae corresponding to the apical (AM) and basolateral (BM) membranes. * p > 0.05; ** p < 0.01; *** p < 0.001 and **** p < 0.0001 relative to corresponding data at 4 pdf and 6 dpf (Student t test).

It appeared that membrane order of epithelial cells treated with cholesterol- mβCD had recovered more than in larvae treated with 7KC and mβCD, particularly in the case of intracellular membranes, which demonstrated an almost complete recovery

(Figure 4-4D). Comparison between GP values of apical, basolateral and intracellular membranes of liver epithelial cells at 4 dpf and 6 dpf showed a significant increase confirming that membrane order had recovered in the liver (Figure 4-4D). To confirm the membrane order recovery at 6 dpf stage, an additional comparison was done between the differences of GP values of sterol-treated larvae at 4 dpf against their control versus differences of GP values of post-treated larvae at 6 dpf larvae against their control. The Δ GP values of apical membranes of gut epithelial cells post-treated with 7KC, cholesterol-mβCD and mβCD at 6 dpf were lower than in those sterol-treated at 4 dpf (p = 0.0003, p = 0.0006, and p = 0.0023 respectively), while Δ GP values of 126

basolateral membranes of gut epithelial cells post-treated with 7KC, cholesterol-mβCD and mβCD at 6 dpf were lower than in those sterol-treated at 4 dpf (p = 0.0247, p =

0.0002, and p = 0.0018 respectively) (Figure 4-5). In kidney epithelial cells post-treated with 7KC, cholesterol-mβCD, and mβCD the Δ GP values of apical membranes at 6 dpf were lower than in those sterol-treated at 4 dpf (p < 0.0001). Δ GP values of basolateral membranes of kidney epithelial cells post-treated with 7KC, cholesterol-mβCD and mβCD at 6 dpf were lower than in those sterol-treated at 4 dpf (p = 0.0008, p = 0.0064, and p < 0.0001, respectively) (Figure 4-5). The Δ GP values of apical membranes of liver epithelial cells post-treated with 7KC, cholesterol-mβCD and mβCD at 6 dpf were significantly less than those in sterol-treated cells at 4 dpf (p < 0.0001, p = 0.0079, and p < 0.0001, respectively), while Δ GP values of basolateral membranes of liver epithelial cells post-treated with 7KC, cholesterol-mβCD and mβCD at 6 dpf were slightly lower than those in sterol-treated at 4 dpf (p = 0.0011, p = 0.0301, and p =

0.0009, respectively) (Figure 4-5). In summary, the data showed that Δ GP of apical and basolateral membranes of the gut, kidney and liver of post-treated larvae at 6 dpf were lower than Δ GP of sterol-treated larvae at 4 dpf larvae, and support the membrane order recovery at 6 dpf stage (Figure 4-5).

4.4 Effect of sterol on aPKC apical localization

In order to assess the effect of sterol manipulations and membrane recovery on the localization of polarity proteins in epithelial cells, I immunostained 4 dpf and 6 dpf larvae that were treated with 7KC, cholesterol-mβCD and mβCD at 4 pdf with antibodies against the polarity protein ‘atypical protein kinase C’ (aPKC) and ZO-1 as described in Materials and Methods, Section 2.3.2. 127

To quantify the degree of apical targeting of aPKC, from each group, 10 larvae were imaged with confocal microscopy, and from each larvae, 10 lines were selected in each apical membrane and cytoplasm from each tissue and overall of equal lines = 100.

The fluorescence intensity was measured along the two lines (apical line and cytoplasm line) then apical/cytoplasm ratio of aPKC staining were calculated (Figure 4-6D-F).

128

** ** ** **

** ** **

Figure 4-6: aPKC apical localization in larvae treated with 7KC, cholesterol-mβCD, and mβCD at 4 dpf and after a two-day recovery (6 dpf). (A, B, and C) Zebrafish larvae were left untreated (Control) or treated with 7KC, cholesterol-mβCD or with mβCD at 4 pdf and left to develop to 6 dpf. Larvae were fixed and immuno-stained for aPKC (green, marked by arrows) and ZO-1 (red, marked by arrowheads) while nuclei were stained with DAPI (blue). From confocal images (A, B and C), the ratio of the aPKC intensity of apical (indicated by red line) and cytoplasm (indicated by yellow line) was obtained and calculated (D, E and F) for gut, kidney and liver of 4 and 6 dpf larvae. In A, B and C scale bar = 10 µm. In D, E, and F, data are mean and standard deviations. ns, not significant, p > 0.05; * p < 0.05; ** p < 0.01; **** p < 0.0001 relative to Control (one-way ANOVA with Bonferroni post-test). ** gut lumen.

129

4.4.1 aPKC localization in the gut, kidney and liver of sterol- manipulated larvae at 4 dpf

Figure 4-6A shows gut epithelial cells in 4 dpf larvae after sterol manipulations that were immunostained with aPKC (shown in green), ZO-1 (shown in red) and nuclei stained with DAPI (shown in blue). aPKC localized to the apical membrane and ZO-1 localized to the tight junctions of gut, kidney, and liver epithelial cells. The labelling of

ZO-1 and DAPI showed that the integrity of tight junction and the nuclei shape, respectively, were maintained after sterol manipulations with 7KC, cholesterol-mβCD, and mβCD.

Calculating the ratio of aPKC staining intensity at apical membranes and the cytoplasm showed that treating 4 dpf larvae with 7KC, cholesterol-mβCD and mβCD significantly reduced aPKC apical localization in polarized epithelial cells of the gut, kidney and liver (p < 0.0001) (Figure 4-6D-F) while kidney epithelial cells treated with mβCD showed a lesser but still significant decrease in aPKC apical localization (p =

0.0025) (Figure 4-6E).

The data also showed that cholesterol-mβCD had the highest effect in disrupting aPKC localization in gut epithelial cells. The apical/cytoplasmic ratio of gut cells treated with cholesterol-mβCD was lower than those treated with mβCD (p = 0.0001), but were not different from those treated with 7KC (p = 0.1558). The aPKC apical/cytoplasmic ratio of gut cells treated with 7KC was slightly lower than those treated with mβCD (p = 0.0335) (Figure 4-6D).

The aPKC apical localization in kidney epithelial cells treated with cholesterol- mβCD was slightly lower than in cells treated with mβCD (p = 0.0293), and aPKC

130

apical localization in kidney cells treated with 7KC was similar to that in cells treated with cholesterol-mβCD and mβCD (p = 0.1449 and p = 0.2124, respectively) (Figure 4-

6E).

Again, in liver epithelial cells, the treatment with cholesterol-mβCD reduced aPKC apical localization the most. Significant differences were observed in aPKC apical localization in epithelial cells treated with cholesterol-mβCD and those treated with

7KC (p < 0.0001) (Figure 4-6F). No differences in aPKC apical localization between cholesterol-mβCD- and mβCD-treated larvae was found (p = 0.3634), and the differences between mβCD and 7KC treatments (p = 0.00076) were less than the differences between cholesterol and 7KC (p < 0.0001). This led us to conclude that cholesterol-mβCD had the highest effect on aPKC mislocalization in liver epithelial cells (Figure 4-6F).

4.4.2 aPKC localization in the gut, kidney and liver of post recovery sterol-manipulated larvae at 6 dpf

At 6 dpf stage, aPKC localization in gut epithelial cells showed no significant differences between 7KC and cholesterol-mβCD treated larvae and the Control group (p

= 0.846, p = 0.0932, respectively) while mβCD post-treated larvae showed a slight decrease in aPKC apical localization (p = 0.0055) compared to Control. This indicates that aPKC localization had recovered in epithelial cells in 7KC- and cholesterol-mβCD treated larvae, and to a lesser degree in larvae treated with mβCD (Figure 4-6D).

At 6 dpf, aPKC localization in the kidney epithelial cells treated with 7KC, cholesterol-mβCD and mβCD were similar to Control larvae (p = 0.797, p = 0.0620 and

131

p = 0.3654, respectively) (Figure 4-6E). These data revealed that aPKC apical localization had fully recovered after two days of incubation after sterol manipulations.

After two further days of development, at 6 dpf, only a slight decrease in aPKC apical localization in liver epithelial cells was observed in 7KC- and mβCD-treated larvae (p = 0.0047 and p = 0.0036, respectively), and a minor decrease in cholesterol- mβCD treated larvae (p = 0.0218) compared to Control. This indicates that aPKC localization had recovered post sterol treatment (Figure 4-6F), but not to the same extent as observed in the kidney (Figure 4-6E).

4.5 Conclusion and Discussion

The purpose of first part of this chapter was to manipulate membrane order of epithelial cells of gut, kidney, and liver of 6 dpf zebrafish larvae with different sterol complexes without affecting the cellular structure and then observe the outcomes of this manipulations on membrane order and apical targeting of a polarity protein. The data revealed that all sterol treatments, 7KC, cholesterol-mβCD and mβCD significantly decreased the membrane order of apical, basolateral and intracellular membranes of polarized epithelial cells in the gut, kidney and liver at 6 dpf (Figure 4-2) without affecting the structure and morphology of epithelial cells in the gut, kidney, and liver

(Figure 4-1).

The quantification of GP values confirmed that 7KC, cholesterol-mβCD and mβCD significantly decreased membrane order of the apical, basolateral and intracellular membranes in the three tissues (Figure 4-2A-C). The apical membranes of polarized epithelial cells remained more ordered than the basolateral membrane and intracellular

132

membranes, and basolateral membranes remained more ordered than intracellular membrane under all treatments conditions (Figure 4-4) indicating that the treatments were not specific to certain membranes and tissues. However, some difference could be observed; the treatment with cholesterol-mβCD had almost always the highest effect on decreasing membrane order in the gut, and kidney epithelial cells compared to the other manipulations (Figure 4-3). The reduction of membrane order when treating larvae with

7KC or mβCD was expected and agrees with previous reports in other cells (Massey and Pownall 2005; Rentero et al. 2008). However, it was anticipated that cholesterol- mβCD would increase membrane order of larvae. This was clearly not the case. It is possible that cholesterol dissociated from mβCD during the incubation with larvae and that the free mβCD acted in removing and/or re-distributing cholesterol. Why the treatment with cholesterol-mβCD decreased the membrane order more than mβCD alone is not clear at this point.

Here, 7KC-mβCD, cholesterol-mβCD, and mβCD complexes were found suitable to manipulate the membrane order in living organisms when used at the stated concentrations. Previously, it was found that 7KC showed no adverse impact on cells when used in low concentration (Rentero et al. 2008; Owen et al. 2012) and here the use of 7KC was extended to in vivo studies. Given the difficulties in understanding the mode of action of cholesterol-mβCD, this treatment is not recommended for in vivo studies.

At 6 dpf, Laurdan GP quantification revealed that membrane order of polarized epithelial cells in the gut and liver was decreased more by sterol manipulations than in the kidney. We speculate that this may be related to the gut and liver function.

133

Gut is responsible for digestion and absorption of nutrients as it has absorptive epithelial cells, while hepatocytes, the main polarized epithelial cells in the liver, are essential for the metabolism of a broad range of substances, bile excretion and cholesterol synthesis (Pack et al. 1996). In contrast, the kidney regulates electrolytes, maintains the acid-base balance, and facilitates the reabsorption of pivotal nutrients such as glucose and amino acids. Therefore, gut and liver have more involvement in cholesterol synthesis, metabolism, and absorption than kidney.

As with the membrane manipulations at 6 pdf, 7KC and cholesterol-mβCD treatment and moderate exposure to mβCD significantly reduced membrane order of apical, basolateral and intracellular membranes in the gut, kidney, and liver of 4 dpf larvae. Despite the effect of sterols on the order of the three membranes, the apical membrane remained more ordered than basolateral membrane, and the basolateral membrane remained more ordered than intracellular membranes (Figure 4-4). This also confirmed that the sterol treatments were not specific to certain membranes and tissues.

Although there are only two days difference in our study, the larvae’s organs such as gut and kidney are not yet completely developed at 4 dpf. Therefore, it is possible that the susceptibility toward sterol manipulation in larvae at 4 dpf could be different from that at 6 dpf when organs are completely developed and functional.

After additional two days of development, the effect of sterol manipulations was still noticeable at 6 dpf (Figure 4-4B and D), but all membranes in the three tissues had recovered, and their membrane order was significantly increased compared to the 4 dpf when the sterol manipulations were performed (Figure 4-4D).

134

As was expected, membrane order recovery was most noticeable in the kidney as its membrane order was less affected at 4 dpf than that of the gut and liver. The rapid recovery of membrane order in all tissues after two days of further development confirmed the regenerative ability of the zebrafish, as it can restore function after an injury to several organs such as in the kidney (Huang et al. 2013). Besides, zebrafish has been employed as an in vivo model for investigating kidney injury and regeneration

(Zhou et al. 2010).

The recovery data suggest that disrupted membrane order can be restored in polarized epithelial cells of intact living zebrafish and this may also apply to mammals.

Sterol manipulations indicate the fundamental role of cholesterol in the formation and maintenance the rafts domains in polarized epithelial cells, which is consistent with other reports in mammalian cells (Simons and Ikonen, 1997; Incardona and Eaton 2000).

The relationship between membrane order and polarity proteins was the main focus of the second part of this chapter. The same sterol manipulations that decreased membrane order also significantly reduced the apical targeting of aPKC directly after the treatment at 4 dpf (Figure 4-6). It was noticed that aPKC immunostaining was specific for the apical membrane in gut, kidney, and liver, even after the sterol manipulations (Figure 4-6A-C). Even though all treatments significantly decreased the localization of aPKC, it seems that cholesterol-mβCD treatment had the strongest effect on the apical localization of aPKC in the gut, kidney, and liver. As cholesterol-mβCD decreased the aPKC localization more than mβCD, this indicates that either high level of cholesterol affected the apical distribution of aPKC, or as we mentioned previously, that cholesterol may dissociated from mβCD during the incubation with larvae and that

135

the free mβCD acted in depleting and/or re-distributing cholesterol that then disrupted aPKC localization. As cholesterol has a fundamental role in membrane order of epithelial cells and in membrane protein functions, it seems likely that it also has an important role in the localization and distribution of polarity proteins in the cell membrane.

It was noticed that the apical targeting of aPKC completely recovered in the kidney epithelial cells after sterol manipulations (Figure 4-6E). Therefore, the full recovery of aPKC localization in the kidney occurred in parallel to the membrane order recovery since the membrane order in the kidney also fully recovered after sterol manipulations.

Hence there is a strong correlation between high membrane order and apical localization of aPKC after acute sterol manipulations and post recovery (Figure 4-4, Figure 4-6).

In conclusion, the maximum recovery in membrane order observed in the kidney also resulted in the maximum recovery of aPKC apical targeting in this tissue, proposing a functional association between the two processes. The next chapter aims to confirm this relationship by knocking down the expression of two of polarity proteins,

Par3 and Crb3a, and measuring the knockdown effect on membrane order of polarized epithelial cells in the gut, kidney, and liver.

136

Chapter 5: Reduced expression of polarity proteins and membrane order

5.1 Introduction

Gene knockdown technology is considered one of most powerful approaches to study the function of a specific protein by preventing the expression of the targeted protein. This technique has been applied to several model organisms, such as mice, frogs, and zebrafish by using Morpholino oligos (MOs) (Heasman 2002). Morpholino oligos were first developed by James Summerton in 1985 as a method to block the translation of RNA transcripts in vivo (Partridge et al. 1996; Summerton et al. 1997).

Morpholinos are antisense oligonucleotides that are usually applied as 25 oligomers that bind via complementary nucleic acid base pairing to the RNA of interest. Structurally,

Morpholinos differ from DNA in that the standard nucleic acid bases of morpholinos are bound to morpholine rings as a replacement for deoxyribose rings, and its bases linked together through phosphorodiamidate groups instead of phosphates. Uncharged phosphorodiamidate groups exclude ionization in the typical physiological pH range; thus, morpholinos in cells or organisms are neutrally charged molecules that exhibit a high binding affinity for RNA (Summerton and Weller 1997). This non charged backbone facilitates the movement of MOs in cells by reducing non-specific interactions with other cell components, which make MOs less toxic. Furthermore, MOs are more stable than other antisense oligos because they are resistant to nucleases

(Nasevicius and Ekker 2000; Eisen and Smith 2008).

137

Unlike many antisense structural types such as phosphorothioates and siRNA, morpholinos do not degrade their target RNA molecules, rather they perform "steric blocking” by binding to a target sequence in a RNA strand and simply hinder the molecules that might otherwise interact with the RNA (Summerton 1999). MOs prevent protein production in two different ways, either by splice blocking or by translational blocking. In splice blocking, MOs bind and inhibit pre-mRNA processing via inhibition of the spliceosome components. In translational blocking, MOs bind complementary mRNA sequences within the 5’ untranslated region (UTR) near the translational start site preventing ribosome assembly (Summerton 1999). To assess the level of knockdown, western blots are a reliable way to verify knockdown of the protein of interest. In zebrafish, MOs are usually introduced into the yolks of 1–4 cell-stage embryos and remained effective for several days, producing morphant embryos. Since most of the zebrafish organs are completely formed and functioning within the first 5 days, this technique allows easy investigation of specific protein function in vivo

(Nasevicius and Ekker 2000).

This chapter continues to investigate the relationship between membrane order and polarity proteins. This was done here by knocking down two polarity proteins using morpholinos and observing its effect on the membrane order of epithelial cells and on the localization of aPKC. Par3 and Crb3a are polarity proteins that are highly conserved and belong to the Par and Crumbs complexes, respectively. Both proteins are localized to the apical membrane adjacent to the junctional complex. Due to their significant contribution in the assembly and function of the junctional complex, both Par and

138

Crumbs complexes are responsible for establishing cell polarity in polarized epithelial cells (Bryant and Mostov 2008).

Par3 is expected to have a strong relationship with aPKC as they both exist as a complex that is associated with tight junctions (TJ). It has been found that Par3 directs the aPKC–Par6 complex to certain membrane regions since Par3 frequently arrives at its destination before aPKC and/or Par6 (Harris and Peifer, 2005; Tabuse et al. 1998).

Crumbs (Crb) is assumed to play an important role in stabilizing the subapical localization of the aPKC complex during cellular shape changes. Indeed, Gibson and

Perrimon (2003) have confirmed this by observing that the Crumbs complex stabilizes the apical localization of aPKC without interacting with Par3. On the other hand, the aPKC complex also seems to be required for the stable apical localization of the

Crumbs complex and aPKC is even more critical than Crb in epithelial cell polarity

(Bilder and Perrimon 2003; Tanentzapf and Tepass 2003). The correlation between Par3 and aPKC was investigated by Ishiuchi and Takeichi 2011, and they found a strong relationship between these two proteins during cell polarity. They suggested a cooperative relationship between Willin (an FERM-domain protein) and Par3 in regulating Rho-associated kinases (ROCKs) during cell polarization. Also, it has been shown that depletion of Willin and Par3 completely disrupts aPKC and Par6 localization to the apical junctional complexes (AJCs), and induces an apical contraction caused by the loss of aPKC. The results also revealed that aPKC phosphorylates the ROCKs and suppresses its junctional localization, thus letting cells maintain its normal apical domains (Ishiuchi and Takeichi 2011).

139

In another study, scientists investigated the relationship between aPKC and Par6 in regulating aPKC localization and its catalytic activity. Two different approaches were used to explore this linkage, an in vitro re-formation approach combined with a cultured cell cortical displacement assay to observe aPKC activity under different conditions.

They showed that aPKC was strongly auto-inhibited by two domains in its NH2- terminal regulatory half; a pseudo-substrate motif that occupies the kinase active site, and a C1 domain that supports this activity. Furthermore, aPKC was activated when it interacted with Par6 through PB1 domain heterodimerization. The data revealed the important involvement of Par6 in regulating aPKC activities and localization to promote a high spatial and temporal control of substrate phosphorylation and polarization during cell polarization (Graybill et al. 2012).

In the first part of this chapter, I describe the knockdown of the two polarity proteins, Par3 and Crb3a, by morpholinos and whether this changed the membrane order of epithelial cells in the gut, kidney, and liver and induced any abnormal development on the zebrafish and its internal organs. In the second section, I examined the association between with aPKC by observing the effect of Par3 and Crb3a knockdown on the apical localization of aPKC.

5.2 The effect of Par3 and Crb3a knockdown on membrane order I first examined whether the knockdown of polarity proteins would also result in a loss of membrane order. To do so, I used morpholinos directed against the polarity proteins Par3 (par3 MO) and Crb3a (crb3a MO), which were injected at the 1–4 cell stage and examined these larvae at 4 dpf.

140

In par3 MO injected larvae, 7 of 10 larvae showed a visible phenotype, and 6 of 10 larvae showed a visible phenotype in crb3 MO injected larvae.

5.2.1 Western blot

To confirm the specificity and knockdown effect of the designed morpholinos,

Western blots were performed as described in Section 2.6 in Materials and Methods chapter.

Par3 protein was observed as an approximately 100 kDa molecular weight band in

4 dpf Control larvae and was almost absent from par3 morphants larvae (Figure 5-1A).

Crb3a protein was observed as an approximately 28 kDa molecular weight band in 4 dpf Control larvae and was almost absent from crb3a morphants larvae (Figure 5-1C).

5.2.2 Morpholinos knockdown

Par3 and Crb3a polarity proteins were knocked-down as described in Section 2.5 in the Materials and Methods chapter. The injection of morpholinos resulted in a 84 % and

56 % reduction of the Par3 and Crb3a protein levels respectively at 4 dpf (Figure 5-1), and images are presented of par3 MO and crb3a MO injected larvae (par3 and crb3a morphants) that showed a clear phenotype were selected (Figure 5-2).

141

Figure 5-1: Immunoblot of knockdown (A) Par3 and (B) Crb3a proteins. Immunoblot of protein extracted from 4 dpf larvae that were incubated with either the anti-Par3 polyclonal antibody (A), the anti-Crb3a monoclonal antibody (B), or anti-actin antibody, which was used as a Control to confirm that equivalent amounts of protein were loaded into each lane. The lanes correspond to wild-type larvae (Control), (A) larvae injected with (par3 MO). The 100-kDa Par3 protein are marked with arrow, (B) larvae injected with (crb3a MO). The 28-kDa Crb3a protein are marked with arrow. In (A) and (B) 42- kDa actin proteins are marked with arrows.

Par3 morphants exhibited cardiac edema, a slow heart rate, and delays in their development. Moreover, the yolk occupied most of the gut region with no swim bladder and the par3 morphants larvae were not swimming at 4 dpf. In crb3a morphants, in addition to cardiac edema, an abnormal body curvature (ventral curvature of the spine) was observed, as well as delays in development as in par3 morphants, with no mouth protrusion and no visible swim bladder (Figure 5-2).

142

Figure 5-2: Phenotype of larvae injected with par3 MO and crb3a MO morpholinos. Par3 morphants exhibited cardiac edema, a delay in its development, and no swim bladder. Crb3a morphants also showed cardiac edema in addition to spinal curvature with no a swim bladder.

5.2.3 Laurdan staining and image analysis

To test whether Par3 and Crb3a knockdown altered the membrane order of polarized epithelial cells in the gut, kidney, and liver of 4 dpf zebrafish larvae, a batch of par3 and crb3a morphants, in addition to Control un-injected larvae, were stained with Laurdan as described in Section 2.2.1, fixed, and then imaged to quantify the GP values of epithelial cell membranes of the gut, kidney, and liver as in Section 2.2.2.

143

** **

Figure 5-3: Membrane order in par3 and crb3a morphants larvae. At the 1-4 cell stage, embryos were left un-injected (Control) or injected with morpholinos against Par3 (par3 MO) and Crb3a (crb3a MO) and left to develop to 4 dpf. GP images were obtained from epithelial cells in the gut (A), kidney (B), and liver (C). GP values were obtained from 75 ROI from each apical (AM), basolateral (BM), and intracellular (IM) membranes from 15 larvae. Scale bar = 20 μm. Data shown are the mean and standard deviations. **** p < 0.0001 relative to Control (Student t-test). AM = apical membrane, BM = basolateral membrane, and IM = intracellular membrane. ** gut lumen.

144

5.3 Par3 and Crb3a knockdown results

5.3.1 Membrane order in the gut, kidney, and liver of morphants at 4 dpf

Figure 5-3A–C depicts GP images from the gut, kidney, and liver of Control, par3, and crb3a morphants at the 4 dpf stage. In par3 and crb3a morphants, mild developmental delays by 24 hpf were observed and the gut appeared in 4 dpf morphants as it is at 3 dpf stage in Control larvae. In addition, slight changes in the gut epithelial cells, especially in par3 morphants, were observed after Par3 and Crb3a knockdown

(Figure 5-3A). Kidney epithelial cells were slightly modified after depleting Par3 and

Crb3a proteins as the cells nearly lost their cube-like shape and became relatively longitudinal, particularly in the crb3a morphants (Figure 5-3B). No visible changes were detected in liver epithelial cells, and seemed structurally normal despite the development delay due to Par3 and Crb3a knockdown (Figure 5-3C). Moreover, the GP images indicated a significant decrease in membrane order of apical (red GP color), basolateral (orange-green GP color), and intracellular (green-blue GP color) in the gut, kidney, and liver compared to the Control (Figure 5-3A-C).

GP values of the gut, kidney, and liver were obtained from 15 larvae per group, i.e., for each Control, Par3, and Crb3a group, with a total of ROI = 75 image regions analyzed for each membrane in each tissue. Compared to the Control, the GP values of the apical, basolateral, and intracellular membranes of the gut, kidney, and liver epithelial cells were significantly decreased (p < 0.0001) in par3 and crb3a morphants, which was consistent with the GP coloring. Among the three membranes in the gut, it was noticed that intracellular membrane in both par3 and crb3a morphants was

145

particularly affected by Par3 and Crb3a knockdown as its GP values substantially decreased compared to the Control and also relative to the apical and basolateral membranes (Figure 5-3A).

After knockdown Par3 and Crb3a, the GP values of the apical membrane remained higher than those of the basolateral and intracellular membranes, and the GP values of the basolateral membranes remained higher than the GP values of the intracellular membranes in the gut, kidney, and liver (Figure 5-3).

5.3.4 Tissue comparison

To detect the most affected tissue following polarity protein depletion, we compared the GP values of the three tissues in par3 and crb3a morphants. Statistical analysis revealed that the gut was the most affected tissue in both par3 and crb3a morphants. In par3 and crb3a morphants, the GP values of the apical, basolateral, and intracellular membranes of the epithelial cells of the gut were significantly lower than the GP values of the three epithelial cell membranes of the kidney and liver (p <

0.0001), while no statistically significant differences were observed between the GP values of the kidney and liver in either membranes of both morphants (Figure 5-3), with the exception of the apical membrane of par3 morphants in the kidney that had higher

GP values than those in the liver (p = 0.0005) (Figure 5-3).

5.4 Effect of Par3 and Crb3a knockdown on aPKC localization

After studying the effect of Par3 and Crb3a knockdown on the membrane order of epithelial cells, I investigated the effect of Par3 and Crb3a knockdown on aPKC apical localization. 146

After larvae microinjection with par3 MO and crb3 MO morpholinos in Section 5.2.1, larvae with a visible phenotype (morphants) were fixed and immunostained with antibodies against the polarity protein ‘atypical protein kinase C’ (aPKC) and ZO-1, as described in Section 2.3.3.2 and imaged as in Section 2.3.4.

Figure 5-4: aPKC localization in the epithelial cells of the gut, kidney, and liver in par3 and crb3a morphants larvae. At 1-4 cell stage, embryos were left un-injected (Control) or injected with morpholinos against Par3 (par3 MO) and Crb3a (crb3a MO). They were immunostained for aPKC (green) and ZO-1 (red) and nuclei were stained with DAPI (blue). From the confocal images of (A) gut, (B) kidney, and (C) liver, the apical/cytoplasm ratio for aPKC was determined. In (A), the scale bar is equivalent to 10 µm. In (B), the data are the mean and standard deviations of 100 ROI obtained from 10 larvae. *** p < 0.001; **** p < 0.0001 relative to the Control (one-way ANOVA with Bonferroni post- test). In (A) and (B) ** represent lumen in gut and in kidney; and in (C) * represent liver sinusoid.

147

aPKC localization in the gut

Figure 5-5A shows the gut epithelial cells of 4 dpf larvae labeled with aPKC

(shown in green) at the apical membrane and ZO-1 (shown in red) at the tight junctions of the epithelial cells. In par3 and crb3a morphants, despite the slight development delays, ZO-1 labeling labeled tight junctions even after Par3 and Crb3a depletion. The shape of the nucleus (blue) differed slightly between the Control and par3 and crb3a morphants, which may be due to development delays (Figure 5-3A).

aPKC was localized to the apical membrane but we were unable to determine whether there were any changes in aPKC localization by eye. We further analyzed aPKC targeting as described in Section 4.4.1. The apical/cytoplasm ratio revealed that there was a significant decrease in aPKC apical localization in the epithelial cells of the gut in both par3 and crb3a morphants (p < 0.0001) (Figure 5-4A). aPKC localization in the kidney

In Figure 5-5B, kidney epithelial cells were also stained at the apical regions with aPKC (shown in green), and at the tight junctions with ZO-1 (red). In par3 and crb3a morphants, the integrity of the tight junctions were demonstrated via the ZO-1 labeling after Par3 and Crb3a knockdown, while the appearance of the nucleus (blue) differed to some extent in par3 and crb3a morphants as they became slightly elongated, similar to the slight cells elongation which was observed in GP images of Laurdan staining

(Figure 5-3B). The apical/cytoplasm ratio revealed that there was a significant decrease in aPKC apical localization in par3 and crb3a morphants (p < 0.0001) in the epithelial cells of the kidney (Figure 5-4B).

148

aPKC localization in the liver

Figure 5-5C shows the aPKC (green) and ZO-1 (red) labeling of the epithelial cells of the liver. In both par3 and crb3a morphants, through observing ZO-1 labeling and the nucleus structure after Par3 and Crb3a knockdown, it seemed that the morphology of liver epithelial cells were not affected by the depletion of the polarity proteins Par3 and Crb3a, as was observed after Laurdan staining of the larvae (Figure 5-3C).

After measuring the apical/cytoplasm ratio, the data revealed that there was a significant decrease in aPKC apical localization in par3 and crb3a morphants (p <

0.0001) in the liver epithelial cells (Figure 5- 4C).

5.5 Conclusion and Discussion

Here, Par3 and Crb3a knockdown in zebrafish seemed to induce developmental delays at 4 dpf with a few changes to its external structure (Figure 5-1), as well as moderate alteration to its internal organs, such as the gut and kidney, as shown in

Figures 5-3A and 5-3B. Cell polarity is required for epithelial tissue development and polarity proteins direct polarity networks and other parts of epithelial differentiation.

Apical proteins form a dynamic network with basolateral polarity proteins to establish apical-basal polarization of epithelial cells (Tepass 2012). Disrupting one of these proteins may affect the development and architecture of epithelial cells as polarity proteins control epithelial cell morphogenesis, and regulate their development and survival. For instance, the reduction in expression of the polarity protein aPKCλ in zebrafish cells produced unusual phenotypes, such as the formation of multiple lumens

149

in the developing intestine, in addition to defects in spindle orientation during progenitor cell division in the neural retina (Horne-Badovinac et al. 2001).

Injecting higher volume of morpholinos may disrupt and arrest the development of zebrafish embryos leading to their death at an early stage; therefore, the injected dosage was adjusted to optimize the phenotype-to-toxicity ratio such that morpholinos injection only caused mild phenotype that kept the embryos alive until the 4 dpf stage. In general, most MOs phenotypes in injected larvae were recognizable during the 2-3 days of development, and the knockdown effects detected up to 5 dpf (Smart et al. 2004; Bill et al. 2008). As morpholinos prevent protein translation by binding to a specific mRNA, it is possible that morpholinos dilution at 4 dpf could allow for partial expression of the par3 and crb3a genes (Figure 5-1).

Different elements may limit the morpholinos knockdown process and its persistence in the cells. Knockdown efficiency of MOs is dependent on the strength of its binding affinity; therefore, diluting MOs may restrict the knockdown process as they are asymmetrically distributed as a result of differential mitotic processes in distinct cellular lineages. Generally, most MOs knockdowns are imperfect; accordingly, small but measurable amounts of protein could be produced. Furthermore, it well known that several genes require only a moderate portion of the normal amount of protein to perform their function efficiently. Hence, regardless of the enduring effects of MOs knockdown on the targeted gene, proteins with long half-life could accumulate and achieve functional larvae with high phenotype thresholds (Eisen and Smith 2008; Bill et al. 2009). In addition, zebrafish have a tremendous regenerative ability, which may explain why we did not observed major changes in the internal organs at 4 dpf.

150

Knockdown of either Par3 or Crb3a polarity proteins resulted in a significant reduction in the membrane order of the apical, basolateral, and intracellular membranes in the gut, kidney, and liver at 4 dpf (Figure 5-3). Here, the significant loss of membrane order in the three tissues due to Par3 and Crb3a knockdown resembled the effect of sterol manipulations, confirming the close relationship between membrane order and polarity proteins (Figure 4-6A). Further, GP quantifications demonstrated that the membrane order of the gut decreased the most among the three examined tissues in par3 and crb3a morphants, implying that these two polarity proteins are expressed at a higher level in the zebrafish gut and/or that their function is more important than in the kidney and liver (Figure 5-3). A relatively higher quantity of ASIP/PAR-3 protein was shown to be needed to assemble cell junctions in the immature epithelia of forestomach and small intestine in mammals (Hirose et al. 2002).

In zebrafish, Crb3a is highly expressed in the otic vesicle and both Crb3a and

Crb3b are expressed in the digestive tract primordium. Generally, crumbs genes in zebrafish demonstrate various expression patterns and these variations in expression suggest that crumbs genes may have a fundamental role in zebrafish development

(Omori and Malicki 2006). In humans and mice, Crb3 is mainly expressed in epithelial tissues and skeletal muscles and is localized to the apical and subapical area of intestinal epithelial cells, indicating that it could play a role in epithelial morphogenesis

(Lemmers et al. 2004).

Similar to the lipid manipulations, the loss of membrane order in morphants was also mimicked in impaired aPKC apical localization in the gut, kidney, and liver of 4 dpf larvae after knocking down two of the main polarity proteins, Par3 and Crb3a

151

(Figure 5-4). This was expected as Par3 and Crb3 have a noticeable linkage with aPKC.

Par3 forms an essential signaling complex with Par6 and aPKC that regulates cell polarity in various cell types of different organisms, ranging from C.elegans through to zebrafish and mammals (Henrique and Schweisguth 2003; Chen and Zhang 2013). It has been proposed that the localization of these proteins is reciprocally dependent as any disruption in any one of those three proteins can disorganize the asymmetric localization of all other PAR proteins (Ebnet et al. 2001; Itoh et al. 2001). Further, several researchers have proposed a direct link between the Crb/Crb3 and the Par complex, which includes aPKC (Sotillos et al. 2004; Walther and Pichaud 2010). An in vitro study done in cell culture and in Drosophila epithelial cells revealed that the cytoplasmic domain of Crumbs can assemble Par6 and aPKC at the plasma membrane and that Crumbs directly interacts with aPKC without any mediators (Kempkens et al.

2006). In addition, in MDCK cells, depletion of Crb3 leads to aPKC mislocalization to the developing apical membrane at the two-cell stage (Schlüter et al. 2009). These findings were confirmed recently when it was showed that direct binding between Crb3 and the aPKC complex in MDCK cells allows the aPKC complex to be completely localized to the apical membrane (Hayase et al. 2013).

In conclusion, the results of this chapter provide evidence for the relationship between membrane order and polarity proteins and the strong association between polarity proteins themselves, Par3 and Crb3a with aPKC, which is consistent with the previous findings of polarity proteins studies.

152

Chapter 6: Reduced expression of Flotillin-1 and Flotillin-2

6.1 Introduction

Flotillin-1/Reggie-2 and Flotillin-2/Reggie-1 are highly conserved proteins that associate with lipid rafts and are thus commonly used as rafts markers (Browman et al.

2007). Flotillin-1 and -2 are linked to the inner leaflet of rafts domains through myristoylation and/or palmitoylation and have the capability to form homo- and hetero- oligomers with each other (Banning et al. 2011). Flotillins/Reggies are ubiquitously expressed and involved in several functions such as endocytosis, phagocytosis and cell signaling (Nichols and Otto 2011) in addition to their important involvement, particularly Flotillin-2, in cell-cell adhesion in human epithelial cells (Kurrle et al.

2013). It has been found in in vivo study that Flotillins/Reggies proteins have fundamental role in regulating axon regeneration in retinal ganglion cells (RGCs), which have an implication in neuronal differentiation, in zebrafish. Flotillin/Reggies role in axon/neurite extension in hippocampal and N2a neurons has been also observed in in vitro study (Munderloh et al. 2009). Furthermore, the importance of Flotillins in cholera toxin intoxication has been observed in zebrafish as these proteins has been found to be involved in cholera toxin trafficking and toxicity, which show the role of

Flotillins in toxin transportation from the plasma membrane to the ER (Saslowsky et al.

2010). However, the precise function of Flotillins/Reggies is still unclear.

In this chapter, I examined the correlation between the expression of Flotillin-1 and

-2 and membrane order, and whether knockdown of Flotillins proteins has an influence on the localization of a polarity protein (aPKC) in epithelial cells. 153

6.2 The effect of Flotillin-1a and Flotillin-2a knockdown on membrane order in zebrafish larvae

Here, I examined whether Flotillin knockdown induces a loss of membrane order of zebrafish epithelial cells. I used morpholinos directed against the lipid rafts proteins

Flot-1a (flot-1a MO) and Flot-2a (flot-2a MO) injected at 1-4 cell stage and resulted in knockdown the Flot-1a and Flot-2a proteins level at 4 dpf larvae (Figure 6-2). In flot-1a

MO injected larvae 6 of 10 larvae showed a visible phenotype, and 6 of 10 larvae showed a visible phenotype in flot-2a MO injected larvae.

6.2.2 Western blot

Western blots were performed to confirm the specificity of the morpholinos knockdown as described in Section 2.6 in Materials and Methods chapter.

Flot-1a protein was detected as an approximately 48 kDa molecular weight band in

4 dpf Control larvae, and was almost missing in flot-1a morphants larvae. Flot-2a protein was detected as an approximately 42 kDa molecular weight band in 4 dpf

Control larvae and was almost absent from flot-2a morphants larvae (Figure 6-1).

6.2.1 Morpholino knockdown

Flot-1a and Flot-2a proteins were knocked-down as described in Section 2.5 in

Materials and Methods chapter. Morpholinos injection decreased Flot-1a and Flot-2a expression by 63 % and 80 %, respectively (Figure 6-1), at 4 dpf. Flot-1a and flot-2a morphants that exhibited detectible phenotypes at 4 dpf were selected (Figure 6-2).

154

Figure 6-1: Immunoblot of knockdown (A) Flot-1a and (B) Flot-2a proteins. Immunoblot of protein extracted from 4 dpf larvae that were incubated with either the anti-Flot-1 monoclonal antibody (A) or anti-Flot-2 polyclonal antibody (B) or anti-actin (A and B) which was used as a control to confirm that equivalent amounts of protein were loaded into each lane. The lanes correspond to wild-type larvae (Control), (A) larvae injected with (flot-1a MO), and (B) larvae injected with (flot-2a MO). The molecular weights are indicated in (A) where the 48-kDa Flot-1a protein bands are marked with an arrow, and in (B) the 42-kDa Flot-2a protein bands are marked with an arrow. In (A) and (B) 42-kDa actin proteins are marked with arrows.

Flot-1a MO and flot-2a MO induced developmental changes to the injected larvae.

Flot-1a morphants showed delay in their development; they seemed to be at 3 dpf stage at 4 dpf because the yolk still exist intact, and slight spinal curvature, abnormal eyes size and no swim bladder was observed. In flot-2a morphants, in addition to cardiac edema, an abnormal body curvature (ventral curvature of the spine) was observed; no mouth protrusion and no visible swim bladder could be detected suggesting a delay in development (Figure 6-2).

155

Figure 6-2: Phenotype of injected larvae with flot-1a MO and flot-2a MO morpholinos. At 4 dpf, Flot-1a morphants exhibited cardiac edema and developmental delay with no swim bladder. Flot-2a morphants also showed cardiac edema in addition to spinal curvature with no swim bladder.

6.2.3 Laurdan staining and image analysis

In order to find out whether Flot-1a and Flot-2a knockdown disrupt the membrane order of polarized epithelial cells in the gut, kidney and liver of 4 dpf zebrafish larvae, a batch of Control un-injected larvae, flot-1a and flot-2a morphants were treated with

Laurdan as in Section 2.2.1, fixed and imaged as in Section 2.2.2.

156

** ** **

Figure 6-3: Membrane order in flot-1a and flot-2a morphants larvae. At 1-4 cell stage, embryos were left un-injected (Control) or injected with morpholinos against Flot-1a (flot-1a MO) and Flot-2a (flot-2a MO) and developed to 4 dpf. GP images were obtained for epithelial cells in gut (A), kidney (B), and liver (C) and GP values obtained from 75 ROI from each apical (AM), basolateral (BM) and intracellular (IM) membranes from 15 larvae. Scale bar = 20 μm. Data are mean and standard deviations. **** p < 0.0001 relative to Control (Student t-test). AM = apical membrane, BM = basolateral membrane, and IM = intracellular membrane. ** gut lumen.

157

6.2.4 Flotillin-1a and Flotillin-2a knockdown results

Membrane order in the gut, kidney and liver

Figure 6-3 shows GP images from the gut of Control, flot-1a, and flot-2a morphants.

Because of the mild development delay, which were produced by Flot-1a and Flot-2 knockdown, the gut appeared like the gut of untreated larvae at 3 dpf. Kidney epithelial cells were not affected after knocking-down Flot-1a and Flot-2a proteins, but the kidney appeared as a 3 days old because of the growth delay (Figure 6-3B). Like in gut and kidney, no changes were detected in liver epithelial cells and they seemed to be structurally normal despite of the developmental delay.

Significant changes in membrane order of apical, basolateral and intracellular membranes in the gut, kidney, and liver were indicated by the GP colors of the apical

(red GP color), basolateral (orange-green GP color), and intracellular (green-blue GP color) compared to Control larvae.GP values of the gut, kidney and liver were acquired from 15 larvae, for each Control, flot-1a and flot-2a morphants, and a total of ROI = 75 image regions for each membrane. As expected, the GP values of apical, basolateral and intracellular membranes of gut epithelial cells of flot-1a and flot-2a morphants were significantly lower than GP values of Control larvae (p < 0.0001), which is consistent with the GP coloring. Flot-1a and Flot-2a knockdown did not affect the differences between three membrane in the three tissues, as the GP values of apical membrane remained higher than those of basolateral and intracellular membranes, and GP values of basolateral membranes remained higher than GP values of intracellular membrane

(Figure 6-3A).

158

6.3 Effect of Flotillin-1a and Flotillin-2a knockdown on aPKC localization

After examining the effect of Flot-1a and Flot-2a knockdown on membrane order of epithelial cells, I investigated the effect of Flot-1a and Flot-2a knockdown on aPKC apical localization. Then, after injected larvae with flot-1a MO and flot-2a MO morpholinos in Section 6.2.1, the morphants with observable phenotype were fixed, sectioned and immunostained with aPKC and ZO-1 as in Chapter 2, Section 2.3.2.

Figure 6-3A-C shows the confocal images of the gut, kidney and liver of 4 dpf larvae that labeled with aPKC (shown in green) at the apical membrane, and ZO-1

(shown in red) at the tight junctions of the epithelial cells. In the gut, kidney, and liver flot-1a and flot-2a morphants confocal images showed the tight junctions through ZO-1 staining, and the normal shape of nucleus (blue) that indicated that epithelial cells of these tissues were not affected by the lack of expression of either of the two lipid rafts proteins.

159

** ** **

Figure 6-4: aPKC localization in epithelial cells of the gut, kidney, and liver in flot-1a and flot-2a morphants larvae. At 1-4 cell stage, embryos were left un-injected (Control) or injected with morpholinos against Flot-1a (flot-1a MO) and Flot-2a (flot-2a MO) were immunostained for aPKC (green) and ZO-1 (red) and nuclei were stained with DAPI (blue). From confocal images of (A) gut, (B) kidney, and (C) liver, the apical/cytoplasm ratio for aPKC was determined. In A, scale bar = 10 µm. In B, data are mean and standard deviations of 100 ROI obtained from 10 larvae. ns indicates p > 0.05 relative to Control (one-way ANOVA with Bonferroni post-test). ** represent gut lumen.

aPKC targeting was analyzed as in Section 4.4, and the apical/cytoplasm ratio revealed that there were no significant differences in aPKC apical localization in gut epithelial cells in both flot-1a and flot-2a morphants (p = 0.2388 and p > 0.999 respectively) (Figure 6-3A),

160

as well as in both morphants in kidney (p > 0.999) (Figure 6-4B), and in flot-1a and flot-

2a morphants (p = 0.1973 and p = 0.3812, respectively) in the liver epithelial cells

(Figure 6-4C).

6.4 Conclusion and Discussion

flot-1a MO and flot-2a MO were injected to reduce the expression of Flot-1a and

Flot-2a to almost undetectable levels in zebrafish larvae. This knockdown provoked mild growth delays by approximately 24 hours affecting the outer organs and structure of the larvae while they did not induced any serious changes to the internal organs such gut, kidney and liver.

Zebrafish are characterized by genome duplication for a number of their genes. It has been revealed that zebrafish exhibit up to four different Flotillin/Reggie genes (two

Flotillin-1/ Reggie-2 and two Flotillin-2/Reggie-1 genes) that are ubiquitously expressed during development and adulthood (Málaga-Trillo et al. 2002). However, during development, one of the Flotillin-2/Reggie-1 genes becomes non-functional.

Flotillin-1a/Reggie-2a, Flotillin-1b/Reggie-2b, and Flotillin-2a/Reggie-1a were found to be ubiquitously expressed during the early stages of zebrafish development. Flotillin-

1a/Reggie-2a is expressed in the brain, spinal cord, and neurogenic placodes while

Flotillin-1b/Reggie-2b is expressed in the head mesoderm, neural crest derivatives and along somite borders. Therefore, when Flot-1a and Flot-2a were knocked-down a ventral curvature of the spine was observed in flot-1a and flot-2a morphants (Figure 6-

1). Moreover, Flotillin-2a/Reggie-1a is highly expressed in regions overlapping with the expression pattern of both Flotillin-1/Reggie-2 genes (Von Philipsborn et al. 2005).

161

Flotillin/Reggie have ubiquitous expression, but it seems the reduced effect of the MOs at 4 dpf, which are diluted during further development, in addition to its regeneration ability may be responsible for undetectable changes in gut, kidney, and liver after rafts proteins knockdown.

Flotillins/Reggies have a fundamental role during early development of zebrafish as morpholinos knockdown of Flotillins/Reggies proteins in developing zebrafish induced morphological alterations during gastrulation. A similar development delay was observed in flot-1a and flot-2a morphants. Furthermore, Flotillin-1/Reggie-2 and

Flotillin-2/Reggie-1 are assumed to be mutually dependent on each other as that the protein stability of Flotillin-1/Reggie-2 and the homo and hetero-tetramers structure of

Flotillin/Reggie depends on Flotillin-2/Reggie-1 (Solis et al. 2007). Therefore, it is likely that both genes would show the same phenotype when knockdown.

Flotillin-1a and Flotillin-2a knockdown significantly decreased membrane order of apical, basolateral and intracellular membranes in the gut, kidney, and liver similar to the knockdown of the polarity proteins Par3, and Crb3a (Figure 5-3). These observations are consistent with that Flotillins proteins may have an important involvement in cell function in zebrafish, as its important role has been shown in regulating axon regeneration in retinal ganglion cells (RGCs) in zebrafish (Munderloh et al. 2009). Knockdown of Flotillin-1a and Flotillin-2a induced a decrease in membrane order of the gut epithelial cells more than kidney and liver (Figure 6-2), which also was observed in polarity proteins knockdown (Figure 5-3).

162

Although Flotillin-2 is expressed in different tissues, high expression of Flotillin-2 was recently observed in human enterocytes, which emphasize that Flotillin-2 could have an essential function in intestinal physiology, especially in the colon (Gauss et al. 2013).

Flotillin-1a and Flotillin-2a knockdown significantly reduced the membrane order of epithelial cells in the gut, kidney, and liver at 4 dpf, but it did not disrupt the aPKCs targeting in these cells. This finding suggests that a reduction in membrane order is not sufficient to inhibit the apical targeting of aPKC. Data shown in Chapter 4 suggest that a high membrane order is required for apical localization of aPKC in polarized epithelial cells of gut, kidney, and liver.

163

Chapter 7: Conclusion & Discussion

The aim of the study was to investigate the correlation between membrane order and polarity network in polarized epithelial cells in zebrafish larvae. Laurdan microscopy and immunofluorescence were the main techniques used to characterize and quantify membrane order in live and fixed zebrafish larvae. The data presented in this thesis were obtained by exploiting the optical transparency of Casper zebrafish strain that has advantages over other mutant strains (White et al. 2008). Laurdan microscopy was found to be an effective way to measure membrane order of epithelial cells in vivo.

In this study, Laurdan did not affect the viability and tissue organization (Gaus et al.

2003; Rentero et al. 2008; Owen et al. 2012) and labeling organs with mCherry as previously reported (Owen et al. 2010) was not necessary, which reduced time and expenses. Fixation with PFA that has been frequently used in other studies as a mean to preserve cells (Gaus et al. 2003; Gaus et al. 2005; Rajendran et al. 2009) was also found suitable for zebrafish larvae (Owen et al. 2010; Owen et al. 2012) as it did not affect the difference in membrane order within epithelial cells.

In two previous studies, the apical membrane of epithelial cells was found to be significantly more ordered than the basolateral membrane in cultured cells (Vieira et al.

2006) and in living zebrafish (Owen et al. 2010). This observation was confirmed in

Chapter 3, where apical membranes of epithelial cells in the gut, kidney and liver of zebrafish was found to be more ordered than basolateral and intracellular membranes when membrane order was quantified in vivo, for the first time, during developmental stages from 3-11 dpf. More specifically, membrane order of epithelial cells in these tissues depended on the development stage and peaked at 6 dpf. At this stage, zebrafish 164

organs are completely developed suggesting a link between membrane order and functions of polarized epithelial cells such as apical-basolateral trafficking. That membrane order decreased beyond the 6 pdf stage and larvae start to feed may also point towards a regulation of membrane order by the diet, however, this link requires further investigations.

In Chapter 4, sterol manipulations were used to manipulate membrane order in polarized epithelial cells in vivo. Sterol complexes of 7KC-mβCD, cholesterol-mβCD

(Rentero et al. 2008; Magenau et al. 2011) and mβCD (Christian et al. 1997; Traian et al. 2012) showed no adverse effect on the tissue organization but reduced membrane order of epithelial cells in the gut, kidney, and liver. The differences between apical membrane with basolateral and intracellular membranes were maintained in sterol- modified larvae.

Surprisingly, cholesterol-mβCD reduced membrane order in 6 dpf zebrafish tissues while Rentero et al. 2008 found no such effect in T cells (Rentero et al. 2008). A possible suggestion is that cholesterol may dissociated from mβCD during the incubation with larvae and the resulted free mβCD either removed or re-distributed cholesterol in zebrafish. What we expected is that the cholesterol-mβCD may be dissociated inside zebrafish cells, not only in the water during incubation. For example,

Frijlink et al 1991 previously injected a rat with cholesterol-mβCD in their study and found that this complex dissociated and that cholesterol accumulated in its kidney cells.

There was no clear reason why cholesterol-mβCD gave unexpected results, and this should be subjected to further investigations to understand exactly what happened.

Moreover, the kidney may degrade and utilize the cholesterol that came from

165

cholesterol-mβCD complexes, which would account for the observation that membrane order in kidney epithelial cells was less affected by the sterol complexes than epithelial cells in the gut and liver that are also involved in cholesterol synthesis and metabolism

(Pack et al. 1996). This means that cholesterol-mβCD is not the preferred choice to manipulate membrane order in vivo.

Sterol manipulations at the 4 dpf stage confirmed the observation at the 6 dpf summarized above. However sterol complexes differ in their effect on disrupting membrane order in epithelial cells in the three examined zebrafish tissues, which may be caused by the differences in zebrafish responses toward chemical substances during its development (Praskova et al. 2011; Kovrižnych et al. 2013). It is noteworthy that membrane order in kidney epithelial cells was almost fully recovered after a further 2 days of development, i.e. at 6 dpf, but these cells were also the least affected by the sterol complexes at 4 dpf stage.

The recovery observations were larvae was treated with sterol complexes at 4 pdf and examined at 6 dpf support previous findings regarding the regeneration ability of zebrafish. Here, membrane order and polarity protein localization was restored within two days. The tremendous regeneration ability has been described in different zebrafish tissues such as gut, where it has been found that gut epithelial cells are restored in the same manner as in mammals (Crosnier et al. 2005), kidney, as regeneration of mesonephric nephrons was started after only 2 days of renal injury (Zhou et al. 2010), and liver, where injured hepatocytes started to recover at 5 dpf, then size and structure of liver was restored at 6 dpf and showed a full recovery at 8 dpf (Curado et al. 2010). It would be interesting to be examined in future studies whether the regeneration of tissue,

166

membrane order and polarity networks extends to cholesterol-related diseases in zebrafish. Zebrafish considered a powerful model in lipid research and in studying metabolism because they have all the key organs needed for metabolic control in humans. Various metabolic diseases have been successfully modelled in the zebrafish, therefore, using zebrafish as a model for lipid-related diseases, including atherosclerosis and obesity are increased recently (Stoletov et al. 2009; Seth et al. 2013).

In addition to disrupting membrane order in polarized epithelial cells, 7KC-mβCD, cholesterol-mβCD, and mβCD also disorganized the apical targeting of the polarity protein aPKC evoking the importance of the physical properties of the membranes in epithelial cells for polarity networks. This has been shown in murine adipocytes that were incubated with mβCD; depletion of cholesterol disrupted rafts-associated proteins and prevented aPKC trafficking to the plasma membrane (Kanzaki et al. 2004).

Cholesterol seems to play a fundamental role in the localization and distribution of polarity proteins in the cell membrane of epithelial cells. Cholesterol depletion usually disrupts the integrity of the cell membrane (Prydz and Simons 2001; Ostasov et al.

2007; Xing et al. 2011) and was reported to hinder proteins trafficking from TGN to the apical membrane (Simons and Toomre 2000), which may explain the decrease in aPKC apical targeting observed here.

Simultaneously to the recovery of the membrane order two days after treatment, aPKC apical localization also recovered and the highest recovery was found in the kidney. This strongly supports a relationship between membrane order and polarity networks in polarized epithelial cells. This is considered the first piece of evidence for a functional relationship between polarity proteins and membrane organization in vivo.

167

The relationship between membrane order and polarity networks was confirmed in

Chapter 5 by employing the morpholino approach from the gene technology field

(Heasman 2002). Morpholinos are the best method to knockdown the expression of specific proteins in zebrafish (Nasevicius and Ekker 2000; Bill et al. 2009; Hosen et al.

2013). Here, the reduced expression of polarity proteins, Par3 and Crb3a, decreased membrane order in zebrafish tissues at 4 pdf. This is a novel finding. The data propose that regulating membrane order and polarity networks are interconnected cellular processes and suggest a reciprocal relationship between polarity networks and membrane organization.

In Par3 and Crb3a morphants, the highest reduction in membrane order was observed in the gut epithelial cells, which suggests that these polarity proteins have an important role in gut function and development. In mammals, Par3 has a higher expression in the immature epithelia of forestomach and small intestine (Hirose et al.

2002) while Crb3 localizes to the apical and subapical area of intestinal epithelial cells in human and mice (Lemmers et al. 2004). Crb3a with Crb3b are expressed in the digestive tract primordium of zebrafish (Omori and Malicki 2006). Disruption of aPKC localization by Par3 and Crb3a knockdown confirmed the strong functional relationship between Par and Crumbs complexes and aPKC in establishing and maintaining cell polarity (Sotillos et al. 2004; Kempkens et al. 2006; Schlüter et al. 2009; Walther and

Pichaud 2010; Hayase et al. 2013).

Taking the observations of Chapter 5 together, the data suggest that polarity networks have important implications for the regulation of lipid organization in the cell membrane of polarized epithelial cells. It has been previously shown that polarity

168

complexes (Crumbs, PAR, and Scribble), signaling proteins, including Rho family

GTPases, in addition to the cytoskeleton and the endocytic and exocytic vesicle trafficking pathways together control the distributions of plasma membrane proteins and lipids between apical and basolateral membranes (Nelson 2009). In addition, the asymmetric location of polarity complexes enables the development of membranes with which they associate. It has been found that the Crumbs complex stimulates apical membrane formation while Par complex equally distributes between apical and lateral membranes (Pieczynski and Margolis 2011). After cholesterol is synthesized in the ER and sphingolipids in the Golgi, these membrane lipids are thought to be transported from early endosomes via a Rab-mediated mechanism (Meer 1989). Similar to protein sorting; sphingolipids are sorted to the TGN, and cholesterol is sorted to both the TGN and lysosomes, then they are delivered to their final destination on the plasma membrane of epithelial cells. Their recognition and sorting are not well understood and need to be clarified in more depth (Aït-Slimane and Hoekstra 2002). In summary, different membrane phospholipids are enriched to different membrane domains by the action of polarity networks and membrane trafficking mechanisms (Mellman and

Nelson 2008). Morpholino knockdown was extended to Chapter 6 to explore the relationship between lipid rafts proteins, membrane order and polarity proteins. As we anticipated, knockdown of the rafts proteins Flot-1a and Flot-2a disrupted membrane order in the gut, kidney, and liver, which confirmed the role of these proteins in rafts formation. Flot-1 knockdown was found to disrupt raft formation in oesophageal squamous epithelial cell (Song et al. 2012) and in rat lung epithelial cells that express

Flotillin-1 and -2 (Chintagari et al. 2008).

169

On the other hand, in this study knockdown of lipid raft proteins did not interfere with the apical localization of aPKC in zebrafish larvae. This finding suggests that disruption of membrane order by Flot-1a and Flot-2a knockdown is not sufficient to block apical targeting of aPKC.

As raft proteins, Flot-1a and Flot-2a play a vital role in various cell functions. It has been demonstrated that Flotillins play a key role in cell signaling, endocytosis and organization of the actin and microtubule cytoskeleton (Babuke and Tikkanen 2007;

Nichols and Otto 2011). A strong linkage between Flotillins and endocytosis has been confirmed by different studies (Hansen and Nichols 2009; Aït-Slimane et al. 2009). The first evidence that Flotillins are directly involved in endocytosis is derived from studies on the uptake of the GPI-anchored protein CD59 and the receptor for cholera toxin, the glycosphingolipid GM1. By knocking-down Flotillin 1, and inhibiting dynamin, the internalisation of both CD59 and cholera toxin B subunit (CTxB) was decreased, which strongly affirms the role of Flotillin 1 in the endocytosis of CD59 and CTxB.

Additional data came from the dynamics of Flotillins when they have been observed to bud into the cell during endocytosis and that Flotillins were co-localized with endocytic cargos directly after internalisation (Glebov et al. 2006). In another study, Saslowsky et al. 2010 have revealed that Flotillins are essential for cholera toxin trafficking and toxicity in both cultured cells and in zebrafish (Saslowsky et al. 2010). This was the first evidence of the direct involvement of Flotillins in endocytosis in a model organism.

Flotillins have also been implicated in regulating membrane lipids as they recognize membrane cholesterol and control the uptake of the cholesterol transporter Niemann-

Pick C1-like 1 (NPC1L1) in mice.

170

In addition, it has been observed that during intracellular trafficking, Flotillin localization is regulated by NPC1L1 (Ge et al. 2011). Here, the outcome of these studies showed the important implication of the raft proteins, Flotillins, in endocytosis, intracellular trafficking and in lipid organization.

The outcomes of Chapter 4 propose that a high membrane order is required for apical localization of aPKC in polarized epithelial cells. However, the sterol complexes reduced membrane order more than Flot-1a and Flot-2a knockdown suggesting that there is a threshold in membrane order that is sensed by polarity proteins. We speculate that the mechanism of how these raft proteins function in the cell is responsible for reducing membrane order without affecting aPKC localization. Generally, Flotillins have a role in different functions such as blocking signalling molecules, inhibiting interactions and stopping signalling responses (Allen et al. 2007); however, it has been found that Flotillins are functioning in different ways in the same process. One suggestion for this is that Flotillins sometimes act as a molecular scaffold provider for membrane rafts that behave as a signaling platform or by establishing a space for the delivery of a particular cargo (Stuermer, 2011). Another possibility is that Flotillins behave as a regulator by detecting any changes in membrane properties, such as lipid composition, and responding to that change appropriately. A third possibility proposes that the budding of flotillin microdomains from the plasma membrane describes a specific type of endocytic pathway (Glebov et al. 2006). Therefore, Flot-1a and Flot-2a may affect lipid organization by reducing membrane order in a different mechanism from that of sterol complexes and polarity proteins, which do not affect aPKC localization.

171

It is also possible that Flot-1 expression can compensate for the loss of Flot-2 and vice versa (Solis et al. 2007; Gauss et al. 2013). Hence, knockdown of one variant of

Flotillin may not be sufficient to reduce membrane order below the threshold of the disruption of aPKC apical localization. In addition to the above speculation, the correlation between Flotillins and polarity proteins, particularly aPKC, was not known previously and requires further investigation, especially because both polarity proteins and raft proteins have a strong connection with membrane trafficking and lipid organization.

It is likely that sterol manipulation will affect the membrane order of intracellular membranes as it contains cholesterol, even at low levels. MβCD extracts membrane cholesterol and 7KC induces the fluidity of the membrane, which presumably decreases membrane order of intracellular membranes. On the other hand, the data showed in my study suggest that knocking-down polarity and flotillin proteins may decrease membrane order in two proposed ways; for example by inducing developmental delay, and this can be found in chapter 5, page 142 and (Figure 5-2) page 143, and in chapter 6 page 155 and (Figure 6-2) page 156. This development delay surely affects the membrane order of apical, basolateral and intracellular membranes because it was shown in chapter 3 (Figure 3-8) page 97, that membrane order at 3 dpf is lower than that at 4 dpf, and membrane order at 4 dpf is lower than that at 6 dpf. The other suggestion is that knocking-down polarity and flotillin proteins disrupts membrane trafficking and lipid organization in the cell (Shivas et al. 2010; Ge et al. 2011) which will, as a result, affect the membrane order of apical, basolateral and intracellular membranes.

172

Figure 7-1: Correlation between membrane order and aPKC localization. Differences in aPKC in apical/cytoplasmic ratio of aPKC apical localization (Δ aPKC) between Control and sterol-treated larvae at 4 dpf (green circle), post recovery at 6 dpf (blue circle), and par3 and crb3a morphants (red circle) Δ GP represent differences in GP values of apical membrane in sterol-treated larvae at 4 dpf, 6 dpf, and par3 and crb3a morphants relative to their controls in gut (A), kidney (B), and liver (C). Data are mean and standard deviations.

In conclusion, the data presented here revealed a novel finding between two important cell regulators, membrane order and polarity network. A reduction in membrane order led to mislocalization of the polarity protein aPKC to the apical membrane and reduced expression of the polarity proteins Par3 or Crb3a decreased membrane order in polarized epithelial cells. This correlation was statistically confirmed as the differences in membrane order (Δ GP) and differences in aPKC localization (Δ aPKC apical/cytoplasmic ratio) in sterol-treated larvae at 4 dpf and post recover at 6 dpf, and par3 and crb3a morphants showed a strong correlation between membrane order and polarity network (Figure 7-1). The strongest relationship was found in kidney epithelial cells, which also had a full recovery in membrane order and aPKC apical localization after sterol manipulation. The finding proposes that lipid trafficking to apical and basolateral membranes with the aid of polarity proteins may maintain the differences in membrane order between these membranes.

173

The asymmetry between apical and basolateral membranes is very important for proper function of polarized epithelial cells. It is well known that polarity protein complexes and membrane trafficking are responsible for this asymmetric organization by delivering the proper proteins and lipids to each membrane. Membrane trafficking is a central process in biosynthesis of cells, secretion, and endocytosis. It assists cells to interact with their outside environments and supports cell growth, propagation, and migration. Trafficking also supports intercellular communication and the structure of extracellular matrix through secretion, and the processing of extracellular signals through endocytosis, and through periodic turnover and renewal of cellular organelles.

Membrane trafficking is also fundamental for cells to maintain cellular homeostasis, as well as meeting specific demands during signal perception and transduction (Cheung and de Vries 2008). On the other hand, scientists showed that polarity proteins, Par3,

Par6/PKC-3 and CDC-42, have an important role in regulating intracellular trafficking, including recycling of clathrin-independent cargo. They also found that CDC-42/Cdc42 is enriched on recycling endosomes in mammalian cells and C. elegans, which proposed a direct function in the regulation of transport in endocytosis process (Balklava et al.

2007). Shivas et al. 2010 confirmed the mutual relationship between polarity complexes and membrane trafficking involving a bidirectional impact on their localization and function accordingly, where epithelial polarization implicates not only asymmetry of the cell cortex, but also reorganization of intracellular trafficking pathways (Shivas et al. 2010). For that reason, when cell polarity is not well established, e.g. by loss of one of the polarity proteins, it will certainly affect membrane trafficking that will lead to improper delivery of lipids and proteins to their target membrane. As a result, the membrane order differences in apical and basolateral membranes will be 174

disrupted, leading to many diseases such as polycystic kidney disease, hypercholesterolaemia, and cancer (Mellman and Nelson 2008). Additionally, membrane order may be a trafficking hallmark for apical sorting in epithelial cells.

Finally, the techniques used here may facilitate further studies to better link and gain deeper understanding of the relationship between biology of membranes and the biology of polarized trafficking.

The new viewpoint presented in this study revealed the importance of polarity networks in lipid organization in the cell membrane and confirmed the strong relationship between polarity proteins in the polarized epithelial cells in the gut, kidney, and liver of zebrafish. Further investigations into this relationship will add data towards a complete vision of the functional relationship between polarity proteins and membrane organization. More than one question needs further validation and clarification in this context. Do other zebrafish tissues such as the pancreas, heart, retina, and vascular system show the same relation between membrane order and polarized trafficking? Are other polarity complexes, such as the scribble complex that localizes to the basolateral membrane, regulated by membrane order, and does the scribble complex have a mutual relationship with other polarity proteins such as aPKC?

Implementing super-resolution microscopy will reveal more information about the reciprocal relationship between polarity networks and membrane order. This will significantly improve the ability to resolve the localization of polarity and raft proteins with more accuracy, and it will assist in delivering more accurate data about raft size.

Lipid rafts have been estimated to be around 20 nm in size, which is under the resolution limit, ~ 200 nm, and this makes exploring rafts in living cells and organisms 175

more demanding. Super-resolution microscopy overcome the diffraction limit and allows acquisition of images with higher resolution, nearly 20 nm, which allows imaging of the exact lipid raft size, and makes it possible to distinguish between two fluorescent objects that are closer together than ∼200–300 nm. Accordingly, this will reveal more about raft composition, structure, and function.

Stimulated emission depletion (STED) is one of the super-resolution microscopy techniques that has been used in different biological tissues with higher resolution, around 20 nm or ~ 10 times less than the diffraction limit. During the last few years super-resolution microscopy has been highly improved by the integration of multi-color, live cell and three-dimensional imaging to STED microscopy (Willig et al. 2007;

Vicidomini et al. 2011). STED has been effectively employed in imaging intact living organisms such as C.elegans (Rankin et al. 2011), and zebrafish (Lv et al. 2012).

Selective-plane illumination microscopy (SPIM) was incorporated with STED in imaging zebrafish embryos to promote high-resolution imaging in model organisms. By using this technique, an approximately 60% improvement in axial resolution with lateral resolution enhancements of 11-40% in control samples and zebrafish embryos were obtained. This delivered imaging of live organisms with high resolution and rapid acquisition, > 100 µm deep in the tissue with no need for histological sectioning for fixed organisms or tissues, which decreases additional artifacts (Friedrich et al. 2011).

The STED technique also succeeded in exploring the distribution of ribbon proteins, whose location in 4 dpf zebrafish retina was demanding to analyze using conventional microscopy. This technique facilitated the observation of the expression of RIBEYE and RIM2 proteins along the synaptic ribbon in the photoreceptor cells (Lv et al. 2012).

176

By using STED, high resolutions of 50–100 nm can be accomplished with immunofluorescence labelling, which might resolve the accurate localization of different membrane proteins in zebrafish tissues. Therefore, employing the advantages of STED with a suitable environmental dye will help to expose more about the relationship between membrane organization and the polarity network in different tissues of zebrafish larvae.

177

Bibliography

Aït-Slimane, Tounsia, and Dick Hoekstra. 2002. Sphingolipid Trafficking and Protein Sorting in Epithelial Cells. FEBS Letters. (529) (1) 54–59.

Aït-Slimane, Tounsia, Galmes Romain, Trugnan Germain, and Maurice Michèle. 2009. Basolateral Internalization of GPI-anchored Proteins Occurs via a Clathrin-independent Flotillin-dependent Pathway in Polarized Hepatic Cells. Mol Biol Cell 1; 20(17): 3792– 3800.

Alberts B, Johnson A, Lewis J, et al. 2002. Molecular Biology of the Cell. 4 th Edit. New York: Garland Science.

Alfons T. L. Van Lommel. 2003. From Cells to Organs: A Histology Textbook and Atlas By Alfons T. L. Van Lommel. illustrate. Springer.

Alfsen, A, P Iniguez, E Bouguyon, and M Bomsel. 2001. Secretory IgA Specific for a Conserved Epitope on Gp41 Envelope Glycoprotein Inhibits Epithelial Transcytosis of HIV-1. The Journal of Immunology 166 (10): 6257-6265.

Allen, John A, Robyn A Halverson-Tamboli, and Mark M Rasenick. 2007. Lipid Raft Microdomains and Neurotransmitter Signalling. Nature Reviews Neuroscience 8 (2): 128- 140.

Amsterdam, Adam, and Nancy Hopkins. 2006. Mutagenesis Strategies in Zebrafish for Identifying Genes Involved in Development and Disease. Trends in Genetics 22 (9): 473- 478.

Babuke, Tanja, and Ritva Tikkanen. 2007. Dissecting the Molecular Function of Reggie/flotillin Proteins. European Journal of Cell Biology 86 (9): 525-532.

Bacia, Kirsten, Dag Scherfeld, Nicoletta Kahya, and Petra Schwille. 2004. Fluorescence Correlation Spectroscopy Relates Rafts in Model and Native Membranes. Biophysical Journal 87 (2): 1034–1043.

Bagatolli, L A, E. Gratton, and G. D. Fidelio. 1998. Water dynamic in glycosphingolipids studied by LAURDAN fluorescence. Biophys. Journal 75, 331-341.

Bagatolli, L A, and E Gratton. 2000. Two Photon Fluorescence Microscopy of Coexisting Lipid Domains in Giant Unilamellar Vesicles of Binary Phospholipid Mixtures. Biophysical Journal 78 (1): 290-305.

Balklava, Zita, Saumya Pant, Hanna Fares, and Barth D Grant. 2007. Genome-Wide Analysis Identifies a General Requirement for Polarity Proteins in Endocytic Traffic. Nature Cell Biology 9 (9): 1066–1073.

178

Bandorowicz-Pikuła, J. 2000. Lipid-binding Proteins as Stabilizers of Membrane Microdomains--possible Physiological Significance. Acta Biochimica Polonica 47 (3): 553–564.

Banning, Antje, Ana Tomasovic, and Ritva Tikkanen. 2011. Functional Aspects of Membrane Association of Reggie/Flotillin Proteins. Current Protein and Peptide Science. 12(8):725- 35.

Baumann, C A, V Ribon, M Kanzaki, D C Thurmond, S Mora, S Shigematsu, P E Bickel, J E Pessin, and A R Saltiel. 2000. CAP Defines a Second Signalling Pathway Required for Insulin-stimulated Glucose Transport. Nature 407 (6801): 202-207.

Baumgart, Tobias, Adam T Hammond, Prabuddha Sengupta, Samuel T Hess, David a Holowka, Barbara a Baird, and Watt W Webb. 2007. Large-scale Fluid/fluid Phase Separation of Proteins and Lipids in Giant Plasma Membrane Vesicles. Proceedings of the National Academy of Sciences of the United States of America 104 (9) (February 27): 3165-70.

Berridge, M.J. 1972. The Mode of Action of 5-Hydroxytryptamine. J. Exp. Biol., (56): 311–321.

Bibhas Kar, Sivamani Subbiah. 2013. Zebrafish: An in Vivo Model for the Study of Human Diseases. International Journal of Genetics and Genomics 1 (1): 6–11.

Bilder, David, Markus Schober, and Norbert Perrimon. 2003. Integrated Activity of PDZ Protein Complexes Regulates Epithelial Polarity. Nature Cell Biology 5 (1): 53–58.

Bill, Brent R, Andrew M Petzold, Karl J Clark, Lisa A Schimmenti, and Stephen C Ekker. 2009. A Primer for Morpholino Use in Zebrafish. Zebrafish 6 (1): 69–77.

Bill, Brent R, Darius Balciunas, Joshua A McCarra, Eric D Young, Toua Xiong, Ashley M Spahn, Marta Garcia-Lecea, Vladimir Korzh, Stephen C Ekker, and Lisa A Schimmenti. 2008. Development and Notch Signaling Requirements of the Zebrafish Choroid Plexus. PloS One 3 (9): e3114.

Binder, Wolfgang H, Veronique Barragan, and Fredric M Menger. 2003. Domains and Rafts in Lipid Membranes. Angewandte Chemie (International Ed. in English) 42 (47) (January): 5802-27.

Bloch K. 1991. Cholesterol: evolution of structure and function. Biochemistry of lipids, lipoproteins, and membranes. Vol. 20. Amsterdam: Elsevier, 363-81.

Brand, M, M Granato, C Nusslein-Volhard, Christiane Nusslein-Volhard, and Ralf Dahm. 2002. Keeping and Raising Zebrafish. Oxford University Press.

Breitkopf, Katja, Laura E Nagy, Juliane I Beier, Sebastian Mueller, Honglei Weng, and Steven Dooley. 2009. Current Experimental Perspectives on the Clinical Progression of Alcoholic Liver Disease. Alcoholism: Clinical and Experimental Research 33 (10): 1647–1655.

Bretscher, Mark S, and Sean Munro. 1993. Cholesterol the Golgi Apparatus. Science 261 (5126): 1280-1281.

179

Browman, Duncan T, Maja B Hoegg, and Stephen M Robbins. 2007. The SPFH Domain- containing Proteins: More Than Lipid Raft Markers. Trends in Cell Biology 17 (8) (August): 394-402.

Brown, DA, and E London. 1998. Functions of Lipid Rafts in Biological Membranes. Annual Review of Cell and Developmental Biology 14 (January): 111-36.

Brown DA, and Rose JK. 1992. Sorting of GPI-anchored proteins to glycolipid-enriched membrane subdomains during transport to the apical cell surface. Cell. 7;68(3):533-44.

Brügger, B, R Sandhoff, S Wegehingel, K Gorgas, J Malsam, J B Helms, W D Lehmann, W Nickel, and F T Wieland. 2000. Evidence for Segregation of Sphingomyelin and Cholesterol During Formation of COPI-coated Vesicles. The Journal of Cell Biology 151 (3): 507–18.

Brugman, Sylvia, Kit-Yeng Liu, Dicky Lindenbergh-Kortleve, Janneke N Samsom, Glenn T Furuta, Stephen A Renshaw, Rob Willemsen, and Edward E S Nieuwenhuis. 2009. Oxazolone-induced Enterocolitis in Zebrafish Depends on the Composition of the Intestinal Microbiota. Gastroenterology 137 (5): 1757–1767.e1.

Bryant, David M, and Keith E Mostov. 2008. From Cells to Organs: Building Polarized Tissue. Nature Reviews Molecular Cell Biology 9 (11): 887–901.

Bulgakova, Natalia A, and Elisabeth Knust. 2009. The Crumbs Complex: From Epithelial-cell Polarity to Retinal Degeneration. Journal of Cell Science 122 (Pt 15): 2587–2596.

Calvet JP, Grantham JJ. 2001. The Genetics and Physiology of Polycystic Kidney Disease. Semin Nephrol: 107–123.

Campbell Neil and Jane Reece. 2005. Biology. 7 th Edit. Pearson Education.

Campagnola, Paul J, and Leslie M Loew. 2003. Second-Harmonic Imaging Microscopy for Visualizing Biomolecular Arrays in Cells, Tissues and Organisms. Nature Biotechnology 21 (11): 1356–1360.

Carrasco, Marlene, Maria Joao Amorim, and Paul Digard. 2004. Lipid Raft-dependent Targeting of the Influenza A Virus Nucleoprotein to the Apical Plasma Membrane. Traffic Copenhagen Denmark 5 (12): 979-992.

Cascio, S, and K S Zaret. 1991. Hepatocyte Differentiation Initiates During Endodermal- mesenchymal Interactions Prior to Liver Formation. Development Cambridge England 113 (1): 217–225.

Cerneus, D P, and A van der Ende. 1991. Apical and Basolateral Transferrin Receptors in Polarized BeWo Cells Recycle through Separate Endosomes. The Journal of Cell Biology 114 (6): 1149–1158.

Chen, Huimin, Elaine R Farkas, and Watt W Webb. 2008. In Vivo Applications of Fluorescence Correlation Spectroscopy. Methods in Cell Biology, Volume 89:3–35.

180

Chen, Jia, and Mingjie Zhang. 2013. The Par3/Par6/aPKC Complex and Epithelial Cell Polarity. Experimental Cell Research 319 (10): 1357–64.

Cheung, Alice Y, and Sacco C de Vries. 2008. Membrane Trafficking: Intracellular Highways and Country Roads. Plant Physiology 147 (4) (August): 1451–3.

Chiang, S H, C A Baumann, M Kanzaki, D C Thurmond, R T Watson, C L Neudauer, I G Macara, J E Pessin, and A R Saltiel. 2001. Insulin-stimulated GLUT4 Translocation Requires the CAP-dependent Activation of TC10. Nature 410 (6831) (April): 944-8.

Chichili, Gurunadh R, and William Rodgers. 2007. Clustering of Membrane Raft Proteins by the Actin Cytoskeleton. The Journal of Biological Chemistry 282 (50): 36682–36691.

Chintagari N R, Deming Gou, and Lin Liu. 2008. Knockdown of Flotillin-2 Inhibits Lung Surfactant Secretion by Alveolar Type II Cells. Cell Res 18 (6): 701–703.

Christian, A E, M P Haynes, M C Phillips, and G H Rothblat. 1997. Use of Cyclodextrins for Manipulating Cellular Cholesterol Content. Journal of Lipid Research 38 (11): 2264– 2272.

Chu, Jaime, and Kirsten C Sadler. 2009. New School in Liver Development: Lessons from Zebrafish. Hepatology 50 (5): 1656–1663.

Cooper, Geoffrey M. 2000. The Cell, A Molecular Approach. 2nd Edditi. Sunderland (MA): Sinauer Associates.

Crane, Jonathan M, and Lukas K Tamm. 2004. Role of Cholesterol in the Formation and Nature of Lipid Rafts in Planar and Spherical Model Membranes. Biophysical Journal 86 (5): 2965–2979.

Curado, Silvia, Elke a Ober, Susan Walsh, Paulina Cortes-Hernandez, Heather Verkade, Carla M Koehler, and Didier Y R Stainier. 2010. The Mitochondrial Import Gene tomm22 Is Specifically Required for Hepatocyte Survival and Provides a Liver Regeneration Model. Disease Models & Mechanisms 3 (7-8): 486–95.

David L. Nelson and Michael M. Cox. 2004. Lehninger Principles of Biochemistry. 4 th Editi. W. H. Freeman.

Davis, Mark E, and Marcus E Brewster. 2004. Cyclodextrin-Based Pharmaceutics: Past, Present and Future. Nature Reviews. Drug Discovery 3 (12): 1023–1035. de Almeida, Rodrigo F M, Luis M S Loura, Alexander Fedorov, and Manuel Prieto. 2005. Lipid Rafts Have Different Sizes Depending on Membrane Composition: A Time-Resolved Fluorescence Resonance Energy Transfer Study. Journal of Molecular Biology 346 (4): 1109–1120.

Decaens, Catherine, Marjorie Durand, Brigitte Grosse, and Doris Cassio. 2008. Which in Vitro Models Could Be Best Used to Study Hepatocyte Polarity? Biology of the Cell / Under the Auspices of the European Cell Biology Organization 100 (7) (July): 387–98.

181

Dermine, J F, S Duclos, J Garin, F St-Louis, S Rea, R G Parton, and M Desjardins. 2001. Flotillin-1-enriched Lipid Raft Domains Accumulate on Maturing Phagosomes. The Journal of Biological Chemistry 276 (21): 18507-18512.

Dietrich, C, L A Bagatolli, Z N Volovyk, N L Thompson, M Levi, K Jacobson, and E Gratton. 2001. Lipid Rafts Reconstituted in Model Membranes. Biophysical Journal 80 (3): 1417– 1428.

Dietrich, Christian, Bing Yang, Takahiro Fujiwara, Akihiro Kusumi, and Ken Jacobson. 2002. Relationship of Lipid Rafts to Transient Confinement Zones Detected by Single Particle Tracking. Biophysical Journal 82 (1 Pt 1): 274–284.

Dietrich, Christian, Zoya N Volovyk, Moshe Levi, Nancy L Thompson, and Ken Jacobson. 2001. Partitioning of Thy-1, GM1, and Cross-linked Phospholipid Analogs into Lipid Rafts Reconstituted in Supported Model Membrane Monolayers. Proceedings of the National Academy of Sciences of the United States of America 98 (19): 10642-10647.

Detrich, H W, M W Kieran, F Y Chan, L M Barone, K Yee, J A Rundstadler, S Pratt, D Ransom, and L I Zon. 1995. Intraembryonic Hematopoietic Cell Migration During Vertebrate Development. Proceedings of the National Academy of Sciences of the United States of America 92 (23): 10713–10717.

Detrich, H. William, Monte Westerfield and Leonard I. Zon. 2010. The Zebrafish: Cellular and Developmental Biology. Elsevier.

Dinic, Jelena, Henrik Biverståhl, Lena Mäler, and Ingela Parmryd. 2011. Laurdan and Di-4- ANEPPDHQ Do Not Respond to Membrane-Inserted Peptides and Are Good Probes for Lipid Packing. Biochimica et Biophysica Acta 1808 (1): 298–306.

Dinic, Jelena, Parham Ashrafzadeh, and Ingela Parmryd. 2013. Actin Filaments Attachment at the Plasma Membrane in Live Cells Cause the Formation of Ordered Lipid Domains. Biochimica et Biophysica Acta - Biomembranes 1828 (3): 1102–1111.

Douarin, N.M. Le. 1975. An Experimental Analysis of Liver Development. Med. Biol: 427–55.

Douglass AD, Vale RD. 2005. Single-molecule microscopy reveals plasma membrane microdomains created by protein-protein networks that exclude or trap signaling molecules in T cells. Cell. 17;121(6):937-50.

Drobnik, Wolfgang, Hana Borsukova, Alfred Böttcher, Alexandra Pfeiffer, Gerhard Liebisch, Gerhard J Schütz, Hansgeorg Schindler, and Gerd Schmitz. 2002. Apo AI/ABCA1- dependent and HDL3-mediated Lipid Efflux from Compositionally Distinct Cholesterol- based Microdomains. Traffic Copenhagen Denmark 3 (4): 268–278.

Drummond, I A, A Majumdar, H Hentschel, M Elger, L Solnica-Krezel, A F Schier, S C Neuhauss, et al. 1998. Early Development of the Zebrafish Pronephros and Analysis of Mutations Affecting Pronephric Function. Development Cambridge England 125 (23): 4655–4667.

182

Drummond, Iain A. 2005. Kidney Development and Disease in the Zebrafish. Journal of The American Society of Nephrology 16 (2): 299–304.

Ebnet, Klaus, Atsushi Suzuki, Yosuke Horikoshi, Tomonori Hirose, Maria-Katharina Meyer Zu Brickwedde, Shigeo Ohno, and Dietmar Vestweber. 2001. The Cell Polarity Protein ASIP/PAR-3 Directly Associates with Junctional Adhesion Molecule (JAM). The European Molecular Biology Organization Journal 20 (14): 3738–3748.

Eckert, Gunter P, Urule Igbavboa, Walter E Müller, and W Gibson Wood. 2003. Lipid Rafts of Purified Mouse Brain Synaptosomes Prepared with or Without Detergent Reveal Different Lipid and Protein Domains. Brain Research 962 (1-2): 144-150.

Edeling, Melissa A, Corinne Smith, and David Owen. 2006. Life of a Clathrin Coat: Insights from Clathrin and AP Structures. Nature Reviews Molecular Cell Biology 7 (1): 32–44.

Eduardo Tolosa, Gregor Wenning, Werner Poewe. 2006. The Diagnosis of Parkinson ’ S Disease. The Lancet Neurology 5: 75–86.

Eeman, Marc, and Magali Deleu. 2010. From Biological Membranes to Biomimetic Model Membranes. Biotechnol. Agron. Soc. Environ. 14(4), 719-736

Eisen, Judith S, and James C Smith. 2008. Controlling Morpholino Experiments: Don’t Stop Making Antisense. Development (Cambridge, England) 135 (10): 1735–1743.

Eriksen, Jason L, Zbigniew Wszolek, and Leonard Petrucelli. 2005. Molecular Pathogenesis of Parkinson Disease. Archives of Neurology 62 (3) (March): 353-7.

Fan, Yang-Yi, Lan H Ly, Rola Barhoumi, David N McMurray, and Robert S Chapkin. 2004. Dietary Docosahexaenoic Acid Suppresses T Cell Protein Kinase C Theta Lipid Raft Recruitment and IL-2 Production. The Journal of Immunology 173 (10): 6151-6160.

Fantini, Jacques, Nicolas Garmy, Radhia Mahfoud, and Nouara Yahi. 2004. Lipid Rafts: Structure, Function and Role in HIV, Alzheimer's and Prion Diseases. Expert Reviews in Molecular Medicine 4 (27): 1–22.

Favard, Cyril, Jerome Wenger, Pierre Francois Lenne, and Herve Rigneault. 2011. FCS Diffusion Laws in Two-Phase Lipid Membranes: Determination of Domain Mean Size by Experiments and Monte Carlo Simulations. Biophysical Journal 100 (5): 1242–1251.

Feigenson, Gerald W. 2007. Phase Boundaries and Biological Membranes. Annu. Rev. Biophys. Biomol. Struct. 36: 63–77.

Field, Holly A, Ober, Elke A, Tobias Roeser, Didier Y.R Stainier. 2003. Formation of the digestive system in zebrafish. I. liver morphogenesis. Developmental Biology. (January) (2) 279–290.

Fielding, Christopher J. 2006. Lipid Rafts and Caveolae. San Francisco: Wiely-VCH Verag Gmbh & Co.KGaA.

183

Fisher, Shannon, Elizabeth a Grice, Ryan M Vinton, Seneca L Bessling, Akihiro Urasaki, Koichi Kawakami, and Andrew S McCallion. 2006. Evaluating the Biological Relevance of Putative Enhancers Using Tol2 Transposon-mediated Transgenesis in Zebrafish. Nature Protocols 1 (3) (January): 1297-305.

Fivaz, M. 1999. Landing on Lipid Rafts. Trends in Cell Biology 9 (6) (June): 212-213.

Fleming, Angeleen, Janusz Jankowski, and Paul Goldsmith. 2010. In Vivo Analysis of Gut Function and Disease Changes in a Zebrafish Larvae Model of Inflammatory Bowel Disease: a Feasibility Study. Inflammatory Bowel Diseases 16 (7): 1162–1172.

Flynn, Edward J, Chad M Trent, and John F Rawls. 2009. Ontogeny and Nutritional Control of Adipogenesis in Zebrafish (Danio Rerio). Journal of Lipid Research 50 (8): 1641–1652.

Fortin, Doris L, Matthew D Troyer, Ken Nakamura, Shin-ichiro Kubo, Malcolm D Anthony, and Robert H Edwards. 2004. Lipid Rafts Mediate the Synaptic Localization of Alpha- synuclein. The Journal of Neuroscience 28; 24 (30): 6715-23.

Friedrich, Mike, Qiang Gan, Vladimir Ermolayev, and Gregory S. Harms. 2011. STED-SPIM: Stimulated Emission Depletion Improves Sheet Illumination Microscopy Resolution. Biophysical Journal 100 (8).

Frijlink HW, Eissens AC, Hefting NR, Poelstra K, Lerk CF, Meijer DK. 1991. The effect of parenterally administered cyclodextrins on cholesterol levels in the rat. Pharm Res 8(1):9- 16.

García-Sáez, Ana J, Salvatore Chiantia, and Petra Schwille. 2007. Effect of Line Tension on the Lateral Organization of Lipid Membranes. The Journal of Biological Chemistry 282 (46) (November 16): 33537-44.

Gassama-Diagne, Ama, Wei Yu, Martin Ter Beest, Fernando Martin-Belmonte, Arlinet Kierbel, Joanne Engel, and Keith Mostov. 2006. Phosphatidylinositol-3,4,5-trisphosphate Regulates the Formation of the Basolateral Plasma Membrane in Epithelial Cells. Nature Cell Biology 8 (9): 963–970.

Gault, Christopher R, Lina M Obeid, and Yusuf A Hannun. 2010. An Overview of Sphingolipid Metabolism: From Synthesis to Breakdown. Advances in Experimental Medicine and Biology 688: 1–23.

Gauss, Annika, Inga Buchholz, Alexandra Zahn, Gerd Schmitz, Wolfgang Stremmel, Joachim Fuellekrug, and Robert Ehehalt. 2013. Flotillin-2 Expression in the Human Gut: From a Cell Model to Human Tissue in Health and Inflammatory Bowel Diseases. International Journal of Medical Sciences 10 (10) (January): 1259–70.

Gaus, Katharina, Elena Chklovskaia, Barbara Fazekas de St Groth, Wendy Jessup, and Thomas Harder. 2005. Condensation of the Plasma Membrane at the Site of T Lymphocyte Activation. The Journal of Cell Biology 171 (1): 121–131.

184

Gaus, Katharina, Enrico Gratton, Eleanor P W Kable, Allan S Jones, Ingrid Gelissen, Leonard Kritharides, and Wendy Jessup. 2003. Visualizing Lipid Structure and Raft Domains in Living Cells with Two-photon Microscopy. Proceedings of the National Academy of Sciences of the United States of America 100 (26) (December): 15554-9.

Gaus, Katharina, Leonard Kritharides, Gerd Schmitz, Alfred Boettcher, Wolfgang Drobnik, Thomas Langmann, Carmel M Quinn, Alison Death, Roger T Dean, and Wendy Jessup. 2004. Apolipoprotein A-1 Interaction with Plasma Membrane Lipid Rafts Controls Cholesterol Export from Macrophages. The FASEB Journal Official Publication of the Federation of American Societies for Experimental Biology 18 (3): 574-576.

Ge, Liang, Wei Qi, Li-Juan Wang, Hong-Hua Miao, Yu-Xiu Qu, Bo-Liang Li, and Bao-Liang Song. 2011. Flotillins Play an Essential Role in Niemann-Pick C1-like 1-Mediated Cholesterol Uptake. Proceedings of the National Academy of Sciences of the United States of America 108 (2): 551–556.

Gibson, Matthew C, and Norbert Perrimon. 2003. Apicobasal Polarization: Epithelial Form and Function Current Opinion in Cell Biology 15 (6): 747–752.

Glebov, Oleg O, Nicholas A Bright, and Benjamin J Nichols. 2006. Flotillin-1 Defines a Clathrin-Independent Endocytic Pathway in Mammalian Cells. Nature Cell Biology 8 (1): 46–54.

Goldsmith, Jason R, Jordan L Cocchiaro, John F Rawls, and Christian Jobin. 2013. Glafenine- induced Intestinal Injury in Zebrafish Is Ameliorated by Μ-opioid Signaling via Enhancement of Atf6-dependent Cellular Stress Responses. Disease Models Mechanisms. 6 (1): 146-59.

Gómez-Moutón, Concepción, Jose Luis Abad, Emilia Mira, Rosa Ana Lacalle, Eduard Gallardo, Sonia Jiménez-Baranda, Isabel Illa, Antonio Bernad, Santos Mañes, and Carlos Martínez- A. 2001. Segregation of Leading-edge and Uropod Components into Specific Lipid Rafts During T Cell Polarization. Proceedings of the National Academy of Sciences of the United States of America 98 (17): 9642-9647.

Gowrishankar, Kripa, Subhasri Ghosh, Suvrajit Saha, C. Rumamol, Satyajit Mayor, and Madan Rao. 2012. Active Remodeling of Cortical Actin Regulates Spatiotemporal Organization of Cell Surface Molecules. Cell 149 (6): 1353–1367.

Grandl, Margot, Salim Maa Bared, Gerhard Liebisch, Tobias Werner, Stefan Barlage, and Gerd Schmitz. 2006. E-LDL and Ox-LDL Differentially Regulate Ceramide and Cholesterol Raft Microdomains in Human Macrophages. Cytometry Part A the Journal of the International Society for Analytical Cytology 69 (3): 189–191.

Graybill, C., B. Wee, S. X. Atwood, and K. E. Prehoda. 2012. Partitioning-Defective Protein 6 (Par-6) Activates Atypical Protein Kinase C (aPKC) by Pseudosubstrate Displacement. Journal of Biological Chemistry. 287:21003-21011.

Griffiths, G, and K Simons. 1986. The Trans Golgi Network: Sorting at the Exit Site of the Golgi Complex. Science 234 (4775): 438–443.

185

Crosnier, Cécile, Neil Vargesson, Stephen Gschmeissner, Linda Ariza-McNaughton, Alastair Morrison, and Julian Lewis. 2005. Delta-Notch Signalling Controls Commitment to a Secretory Fate in the Zebrafish Intestine. Development Cambridge England 132 (5): 1093– 1104.

Grosshans, Bianka L, Darinel Ortiz, and Peter Novick. 2006. Rabs and Their Effectors: Achieving Specificity in Membrane Traffic. Proceedings of the National Academy of Sciences of the United States of America 103 (32): 11821–11827.

Gruenberg, Jean. 2003. Lipids in Endocytic Membrane Transport and Sorting. Current Opinion in Cell Biology 15 (4): 382-388.

Grunwald, David Jonah, and Judith S Eisen. 2002. Headwaters of the Zebrafish -- Emergence of a New Model Vertebrate. Nature Reviews Genetics 3 (9): 717-724.

Hagen, J P, and H M McConnell. 1997. Liquid-liquid Immiscibility in Lipid Monolayers. Biochimica Et Biophysica Acta 1329 (1): 7-11.

Hancock, John F. 2006. Lipid Rafts: Contentious Only from Simplistic Standpoints. Nature Reviews Molecular Cell Biology 7 (6): 456-462.

Hansen, Carsten G, and Benjamin J Nichols. 2009. Molecular Mechanisms of Clathrin- Independent Endocytosis. Journal of Cell Science 122 (Pt 11): 1713–1721.

Harder, T, P Scheiffele, P Verkade, and K Simons. 1998. Lipid Domain Structure of the Plasma Membrane Revealed by Patching of Membrane Components. The Journal of Cell Biology 141 (4) (May): 929-42.

Harder, Thomas. 2004. Lipid Raft Domains and Protein Networks in T-cell Receptor Signal Transduction. Current Opinion in Immunology 16 (3): 353-359.

Harris, F M, S K Smith, and J D Bell. 2001. Physical Properties of Erythrocyte Ghosts That Determine Susceptibility to Secretory Phospholipase A2. The Journal of Biological Chemistry 276 (25) (June): 22722-31.

Harris, Faith M, Katrina B Best, and John D Bell. 2002. Use of Laurdan Fluorescence Intensity and Polarization to Distinguish between Changes in Membrane Fluidity and Phospholipid Order. Biochimica et Biophysica Acta 1565 (1) (September): 123–8.

Harris, Tony J C, and Mark Peifer. 2005. The Positioning and Segregation of Apical Cues During Epithelial Polarity Establishment in Drosophila. The Journal of Cell Biology 170 (5): 813–823.

Hattori, Chinatsu, Masashi Asai, Hayato Onishi, Noboru Sasagawa, Yasuhiro Hashimoto, Takaomi C Saido, Kei Maruyama, Shigehiko Mizutani, and Shoichi Ishiura. 2006. BACE1 Interacts with Lipid Raft Proteins. Journal of Neuroscience Research 84 (4): 912-917.

186

Hayase, Junya, Sachiko Kamakura, Yuko Iwakiri, Yoshihiro Yamaguchi, Tomoko Izaki, Takashi Ito, and Hideki Sumimoto. 2013. The WD40 Protein Morg1 Facilitates Par6-aPKC Binding to Crb3 for Apical Identity in Epithelial Cells. The Journal of Cell Biology 200 (5): 635–50.

Heasman, Janet. 2002. Morpholino Oligos: Making Sense of Antisense? Developmental Biology 243 (2): 209–214.

Heberle, Frederick A., Jing Wu, Shih Lin Goh, Robin S. Petruzielo, and Gerald W. Feigenson. 2010. Comparison of Three Ternary Lipid Bilayer Mixtures: FRET and ESR Reveal Nanodomains. Biophysical Journal 99 (10): 3309–3318.

Heerklotz, Heiko, Halina Szadkowska, Thomas Anderson, and Joachim Seelig. 2003. The Sensitivity of Lipid Domains to Small Perturbations Demonstrated by the Effect of Triton. Journal of Molecular Biology 329 (4): 793-799.

Hehnly, Heidi, and Mark Stamnes. 2007. Regulating Cytoskeleton-based Vesicle Motility. FEBS Letters 581 (11): 2112–2118.

Henderson, Robert M, J Michael Edwardson, Nicholas a Geisse, and David E Saslowsky. 2004. Lipid Rafts: Feeling Is Believing. News in Physiological Sciences : an International Journal of Physiology Produced Jointly by the International Union of Physiological Sciences and the American Physiological Society 19 (10) (April): 39–43.

Henrique, Domingos, and François Schweisguth. 2003. Cell Polarity: The Ups and Downs of the Par6/aPKC Complex. Current Opinion in Genetics & Development 13 (4): 341–350.

Henshaw, J B, C A Olsen, A R Farnbach, K H Nielson, and J D Bell. 1998. Definition of the Specific Roles of Lysolecithin and Palmitic Acid in Altering the Susceptibility of Dipalmitoylphosphatidylcholine Bilayers to Phospholipase A2. Biochemistry 37 (30): 10709-10721.

Her, Guor Mour, Yang-Hui Yeh, and Jen-Leih Wu. 2004. Functional Conserved Elements Mediate Intestinal-type Fatty Acid Binding Protein (I-FABP) Expression in the Gut Epithelia of Zebrafish Larvae. Developmental Dynamics : an Official Publication of the American Association of Anatomists 230 (4) (August): 734–42.

Hinzey, Adam H, Michelle a Kline, Sainath R Kotha, Sean M Sliman, Elizabeth S O’Connor Butler, Andrew B Shelton, Travis R Gurney, and Narasimham L Parinandi. 2012. Choice of Cyclodextrin for Cellular Cholesterol Depletion for Vascular Endothelial Cell Lipid Raft Studies: Cell Membrane Alterations, Cytoskeletal Reorganization and Cytotoxicity. Indian Journal of Biochemistry & Biophysics 49 (5): 329–41.

Hirose, Tomonori, Yasushi Izumi, Yoji Nagashima, Yoko Tamai-Nagai, Hidetake Kurihara, Tatsuo Sakai, Yukari Suzuki, et al. 2002. Involvement of ASIP/PAR-3 in the Promotion of Epithelial Tight Junction Formation. Journal of Cell Science 115 (Pt 12): 2485–2495.

Holthuis, J C, T Pomorski, R J Raggers, H Sprong, and G Van Meer. 2001. The Organizing Potential of Sphingolipids in Intracellular Membrane Transport. Physiological Reviews 81 (4): 1689-1723. 187

Hölttä-Vuori, Maarit, Riikka-Liisa Uronen, Jarmila Repakova, Emppu Salonen, Ilpo Vattulainen, Pertti Panula, Zaiguo Li, Robert Bittman, and Elina Ikonen. 2008. BODIPY-cholesterol: a New Tool to Visualize Sterol Trafficking in Living Cells and Organisms. Traffic (Copenhagen, Denmark) 9 (11) (November): 1839-49.

Hölttä-Vuori, Maarit, Veijo T. V. Salo, Lena Nyberg, Christian Brackmann, Annika Enejder, Pertti Panula and Elina Ikonen. 2010. Zebrafish: gaining popularity in lipid research. Biochem. J. 429 (235–242).

Honigmann, Alf, Veronika Mueller, Stefan W. Hell, and Christian Eggeling. 2013. STED Microscopy Detects and Quantifies Liquid Phase Separation in Lipid Membranes Using a New Far-Red Emitting Fluorescent Phosphoglycerolipid Analogue. Faraday Discussions. 161:77-89.

Hooper, N M. 1999. Detergent-insoluble Glycosphingolipid/cholesterol-rich Membrane Domains, Lipid Rafts and Caveolae (review). Molecular Membrane Biology 16 (2): 145- 156.

Horejsí, Václav. 2004. Transmembrane Adaptor Proteins in Membrane Microdomains: Important Regulators of Immunoreceptor Signaling. Immunology Letters 92 (1-2): 43–49.

Horne-Badovinac, S, D Lin, S Waldron, M Schwarz, G Mbamalu, T Pawson, Y Jan, D Y Stainier, and S Abdelilah-Seyfried. 2001. Positional Cloning of Heart and Soul Reveals Multiple Roles for PKC Lambda in Zebrafish Organogenesis. Current Biology : CB 11 (19): 1492–1502.

Hosen, Mohammad Jakir, Olivier M Vanakker, Andy Willaert, Ann Huysseune, Paul Coucke, and Anne De Paepe. 2013. Zebrafish Models for Ectopic Mineralization Disorders: Practical Issues from Morpholino Design to Post-Injection Observations. Frontiers in Genetics 4 (May): 74.

Howarth, Deanna L, Chunyue Yin, Karen Yeh, and Kirsten C Sadler. 2013. Defining Hepatic Dysfunction Parameters in Two Models of Fatty Liver Disease in Zebrafish Larvae. Zebrafish 10 (2) (June): 199–210.

Howarth, Deanna L, Mike Passeri, and Kirsten C Sadler. 2011. Drinks Like a Fish: Using Zebrafish to Understand Alcoholic Liver Disease. Alcoholism: Clinical and Experimental Research 35 (5): 826–829.

Huang, Jianmin, Mary McKee, Hong D Huang, Alice Xiang, Alan J Davidson, and Hua A J Lu. 2013. A Zebrafish Model of Conditional Targeted Podocyte Ablation and Regeneration. Kidney International 83 (6): 1193–200.

Huang, Shih Hao, Chung Der Hsiao, Dar Shong Lin, Cho Yen Chow, Chia Jen Chang, and Ian Liau. 2011. Imaging of Zebrafish In Vivo with Second-Harmonic Generation Reveals Shortened Sarcomeres Associated with Myopathy Induced by Statin. PLoS ONE 6 (9).

Hung, T J, and K J Kemphues. 1999. PAR-6 Is a Conserved PDZ Domain-containing Protein That Colocalizes with PAR-3 in Caenorhabditis Elegans Embryos. Development Cambridge England 126 (1): 127–135. 188

Hurd, Toby W, Lin Gao, Michael H Roh, Ian G Macara, and Ben Margolis. 2003. Direct Interaction of Two Polarity Complexes Implicated in Epithelial Tight Junction Assembly. Nature Cell Biology 5 (2): 137–42.

Hutterer, A., Betschinger, J., Petronczki, M., Knoblich, J.A. 2004. Sequential roles of Cdc42, Par-6, aPKC, and Lgl in the establishment of epithelial polarity during Drosophila embryogenesis. Dev. Cell 6(6): 845--854.

Ikonen, E. 2001. Roles of Lipid Rafts in Membrane Transport. Current Opinion in Cell Biology 13 (4): 470-477.

Incardona, J P, and S Eaton. 2000. Cholesterol in Signal Transduction. Current Opinion in Cell Biology 12 (2): 193–203.

Ipsen, J H, O G Mouritsen, and M J Zuckermann. 1989. Theory of Thermal Anomalies in the Specific Heat of Lipid Bilayers Containing Cholesterol. Biophysical Journal 56 (4) (October): 661-7.

Ishiuchi, Takashi, and Masatoshi Takeichi. 2011. Willin and Par3 Cooperatively Regulate Epithelial Apical Constriction through aPKC-Mediated ROCK Phosphorylation. Nature Cell Biology 13 (7): 860–866.

Itoh, M, H Sasaki, M Furuse, H Ozaki, T Kita, and S Tsukita. 2001. Junctional Adhesion Molecule (JAM) Binds to PAR-3: A Possible Mechanism for the Recruitment of PAR-3 to Tight Junctions. The Journal of Cell Biology 154 (3): 491–497.

Izaddoost, Shayan, Sang-Chul Nam, Manzoor A Bhat, Hugo J Bellen, and Kwang-Wook Choi. 2002. Drosophila Crumbs Is a Positional Cue in Photoreceptor Adherens Junctions and Rhabdomeres. Nature 416 (6877): 178–183.

Izumi, Yasushi, Tomonori Hirose, Yoko Tamai, Syu-ichi Hirai, Yoji Nagashima, Toyoshi Fujimoto, Yo Tabuse, Kenneth J Kemphues, and Shigeo Ohno. 1998. An Atypical PKC Directly Associates and Colocalizes at the Epithelial Tight Junction with ASIP, a Mammalian Homologue of Caenorhabditis Elegans Polarity Protein PAR-3. The Journal of Cell Biology 143 (1): 95–106.

Jacobowitz DM and Kallaral AT. 2004. Flotillin-1 in the Substantia Nigra of the Parkinson Brain and a Predominant Localization in Catecholaminergic Nerves in the Rat Brain. Neurotox Res. 6(4):245-57

Janes, Peter W, Steven C Ley, and Anthony I Magee. 1999. Aggregation of Lipid Rafts Accompanies Signaling via the T Cell Antigen Receptor. The Journal of Cell Biology 147 (2): 447-461.

Jayachandran, Pradeepa, Elim Hong, and Rachel Brewster. 2010. Labeling and Imaging Cells in the Zebrafish Hindbrain. Journal of Visualized Experiments JoVE 25;(41).

189

Jin, Hao, Raman Sood, Jin Xu, Fenghua Zhen, Milton A English, P Paul Liu, and Zilong Wen. 2009. Definitive Hematopoietic Stem/progenitor Cells Manifest Distinct Differentiation Output in the Zebrafish VDA and PBI. Development Cambridge England 136 (4): 647– 654.

Jin, Lei, Andrew C Millard, Joseph P Wuskell, Xuemei Dong, Dianqing Wu, Heather a Clark, and Leslie M Loew. 2006. Characterization and Application of a New Optical Probe for Membrane Lipid Domains. Biophysical Journal 90 (7) (April): 2563-75.

Johnson, Corey M., and William Rodgers. 2008. Spatial Segregation of Phosphatidylinositol 4,5-Bisphosphate (PIP2) Signaling in Immune Cell Functions. Immunol Endocr Metab Agents Med Chem . 8 (4): 349-357.

Kabouridis, Panagiotis S, Julia Janzen, Anthony L Magee, and Steven C Ley. 2000. Cholesterol Depletion Disrupts Lipid Rafts and Modulates the Activity of Multiple Signaling Pathways in T Lymphocytes. European Journal of Immunology 30 (3): 954–963.

Kai Simons, Robert Ehehalt. 2002. Cholesterol, lipid rafts, and disease. J Clin Invest.110(5):597–603

Kamiguchi, Hiroyuki. 2006. The Region-specific Activities of Lipid Rafts During Axon Growth and Guidance. Journal of Neurochemistry 98 (2): 330-335.

Kanzaki, Makoto, Silvia Mora, Joseph B Hwang, Alan R Saltiel, and Jeffrey E Pessin. 2004. Atypical Protein Kinase C (PKCzeta/lambda) Is a Convergent Downstream Target of the Insulin-Stimulated Phosphatidylinositol 3-Kinase and TC10 Signaling Pathways. The Journal of Cell Biology 164 (2): 279–290.

Karlsson, J, J Von Hofsten, and P E Olsson. 2001. Generating Transparent Zebrafish: a Refined Method to Improve Detection of Gene Expression During Embryonic Development. Marine Biotechnology 3 (6): 522–527.

Karnovsky, M J, a M Kleinfeld, R L Hoover, and R D Klausner. 1982. The Concept of Lipid Domains in Membranes. The Journal of Cell Biology 94 (1) (July): 1-6.

Keller, P, and K Simons. 1997. Post-Golgi Biosynthetic Trafficking. Journal of Cell Science 110 ( Pt 2 (Pt 24): 3001–3009.

Keller, S L, T G Anderson, and H M McConnell. 2000. Miscibility Critical Pressures in Monolayers of Ternary Lipid Mixtures. Biophysical Journal 79 (4): 2033-2042.

Kempkens, Ozlem, Emmanuelle Médina, Gregorio Fernandez-Ballester, Susann Ozüyaman, André Le Bivic, Luis Serrano, and Elisabeth Knust. 2006. Computer Modelling in Combination with in Vitro Studies Reveals Similar Binding Affinities of Drosophila Crumbs for the PDZ Domains of Stardust and DmPar-6. European Journal of Cell Biology 85 (8): 753–767.

Kenworthy, A K, and M Edidin. 1998. Distribution of a Glycosylphosphatidylinositol-Anchored Protein at the Apical Surface of MDCK Cells Examined at a Resolution of <100 A Using Imaging Fluorescence Resonance Energy Transfer. The Journal of Cell Biology 142 (1): 69–84. 190

Kenworthy, A K, N Petranova, and M Edidin. 2000. High-Resolution FRET Microscopy of Cholera Toxin B-Subunit and GPI-Anchored Proteins in Cell Plasma Membranes. Molecular Biology of the Cell 11 (5): 1645–1655.

Kimmel, C B, W W Ballard, S R Kimmel, B Ullmann, and T F Schilling. 1995. Stages of Embryonic Development of the Zebrafish. Developmental Dynamics an Official Publication of the American Association of Anatomists 203 (3): 253-310.

Kim, Hwan Myung, Hyo-Jung Choo, Soon-Young Jung, Young-Gyu Ko, Won-Hwa Park, Seung-Joon Jeon, Chul Hoon Kim, Taiha Joo, and Bong Rae Cho. 2007. A Two-Photon Fluorescent Probe for Lipid Raft Imaging: C-Laurdan. Chembiochem : A European Journal of Chemical Biology 8 (5) (March): 553–9.

Klausner, Richard D, Alan M Kleinfeld, Richard L Hoover, and Morris J Karnovskys. 1980. Evidence derived from structural perturbations induced by free fatty acids and lifetime heterogeneity analysis. J BIOL CHEM 255 (4): 1286-1295.

Klebes, A, and E Knust. 2000. A Conserved Motif in Crumbs Is Required for E-cadherin Localisation and Zonula Adherens Formation in Drosophila. Current Biology 10 (2): 76– 85.

Klein, U, G Gimpl, and F Fahrenholz. 1995. Alteration of the Myometrial Plasma Membrane Cholesterol Content with Beta-Cyclodextrin Modulates the Binding Affinity of the Oxytocin Receptor. Biochemistry 34 (42): 13784–13793.

Klemm, Robin W, Christer S Ejsing, Michal a Surma, Hermann-Josef Kaiser, Mathias J Gerl, Julio L Sampaio, Quentin de Robillard, et al. 2009. Segregation of Sphingolipids and Sterols during Formation of Secretory Vesicles at the Trans-Golgi Network. The Journal of Cell Biology 185 (4) (May): 601–12.

Korzeniowski, Marek, Katarzyna Kwiatkowska, and Andrzej Sobota. 2003. Insights into the Association of FcgammaRII and TCR with Detergent-resistant Membrane Domains: Isolation of the Domains in Detergent-free Density Gradients Facilitates Membrane Fragment Reconstitution. Biochemistry 42 (18): 5358-5367.

Kovrižnych, Jevgenij a, Ružena Sotníková, Dagmar Zeljenková, Eva Rollerová, Elena Szabová, and Soňa Wimmerová. 2013. Acute Toxicity of 31 Different Nanoparticles to Zebrafish (Danio Rerio) Tested in Adulthood and in Early Life Stages - Comparative Study. Interdisciplinary Toxicology 6 (2): 67–73.

Kurrle, Nina, Frauke Völlner, Rüdiger Eming, Michael Hertl, Antje Banning, and Ritva Tikkanen. 2013. Flotillins Directly Interact with Γ-Catenin and Regulate Epithelial Cell- Cell Adhesion. PloS One 8 (12): e84393.

Kusumi, Akihiro, Hiroshi Ike, Chieko Nakada, Kotono Murase, and Takahiro Fujiwara. 2005. Single-Molecule Tracking of Membrane Molecules: Plasma Membrane Compartmentalization and Dynamic Assembly of Raft-Philic Signaling Molecules. Seminars in Immunology 17 (1): 3–21.

191

Kwik, Jeanne, Sarah Boyle, David Fooksman, Leonid Margolis, Michael P Sheetz, and Michael Edidin. 2003. Membrane Cholesterol, Lateral Mobility, and the Phosphatidylinositol 4,5- bisphosphate-dependent Organization of Cell Actin. Proceedings of the National Academy of Sciences of the United States of America 100 (24): 13964-13969.

Lang, D M, S Lommel, M Jung, R Ankerhold, B Petrausch, U Laessing, M F Wiechers, H Plattner, and C A Stuermer. 1998. Identification of Reggie-1 and Reggie-2 as Plasmamembrane-associated Proteins Which Cocluster with Activated GPI-anchored Cell Adhesion Molecules in Non-caveolar Micropatches in Neurons. Journal of Neurobiology 37 (4): 502-523.

Langheinrich, Ulrike. 2003. Zebrafish: a New Model on the Pharmaceutical Catwalk. BioEssays News and Reviews in Molecular Cellular and Developmental Biology 25 (9): 904–912.

Langhorst, M F, a Reuter, and C a O Stuermer. 2005a. Scaffolding Microdomains and Beyond: The Function of Reggie/flotillin Proteins. Cellular and Molecular Life Sciences : CMLS 62 (19-20) (October): 2228–40.

Langhorst, M F, a Reuter, and C a O Stuermer. 2005b. Scaffolding Microdomains and Beyond: The Function of Reggie/flotillin Proteins. Cellular and Molecular Life Sciences : CMLS 62 (19-20) (October): 2228–40.

LaRocca, Timothy J., Priyadarshini Pathak, Salvatore Chiantia, Alvaro Toledo, John R. Silvius, Jorge L. Benach, and Erwin London. 2013. Proving Lipid Rafts Exist: Membrane Domains in the Prokaryote Borrelia Burgdorferi Have the Same Properties as Eukaryotic Lipid Rafts. PLoS Pathogens 9 (5).

Le Douarin, N.M., 1975. An experimental analysis of liver development. Med. Biol. 53, 427– 455.

Lee, Marcus C S, Elizabeth A Miller, Jonathan Goldberg, Lelio Orci, and Randy Schekman. 2004. Bi-directional Protein Transport Between the ER and Golgi. Annual Review of Cell and Developmental Biology 20 (1): 87–123.

Lemmers, Céline, Didier Michel, Lydie Lane-Guermonprez, Marie-Hélène Delgrossi, Emmanuelle Médina, Jean-Pierre Arsanto, and André Le Bivic. 2004. CRB3 Binds Directly to Par6 and Regulates the Morphogenesis of the Tight Junctions in Mammalian Epithelial Cells. Molecular Biology of the Cell 15 (3): 1324–1333.

Lenne, Pierre-François, Laure Wawrezinieck, Fabien Conchonaud, Olivier Wurtz, Annie Boned, Xiao-Jun Guo, Hervé Rigneault, Hai-Tao He, and Didier Marguet. 2006. Dynamic Molecular Confinement in the Plasma Membrane by Microdomains and the Cytoskeleton Meshwork. The European Molecular Biology Organization Journal 25 (14): 3245-3256.

Levental, Ilya, Michal Grzybek, and Kai Simons. 2011. Raft Domains of Variable Properties and Compositions in Plasma Membrane Vesicles. Proceedings of the National Academy of Sciences of the United States of America 108 (28): 11411–11416.

192

Levental, Ilya, Daniel Lingwood, Michal Grzybek, Unal Coskun, and Kai Simons. 2010. Palmitoylation Regulates Raft Affinity for the Majority of Integral Raft Proteins. Proceedings of the National Academy of Sciences of the United States of America 107 (51): 22050–22054.

Li, I-Chen, Chein-Tso Chan, Yu-Fen Lu, Yi-Ting Wu, Yi-Chung Chen, Guo-Bin Li, Che-Yi Lin, and Sheng-Ping L Hwang. 2011. Zebrafish Krüppel-Like Factor 4a Represses Intestinal Cell Proliferation and Promotes Differentiation of Intestinal Cell Lineages. Edited by Vincent Laudet. PLoS ONE 6 (6): 14.

Lingwood, Daniel, and Kai Simons. 2010. Lipid Rafts as a Membrane-organizing Principle. Science (New York, N.Y.) 327 (5961) (January): 46-50.

Li, Qiurong, Meng Wang, Li Tan, Chang Wang, Jian Ma, Ning Li, Yousheng Li, Guowang Xu, and Jieshou Li. 2005. Docosahexaenoic Acid Changes Lipid Composition and Interleukin- 2 Receptor Signaling in Membrane Rafts. Journal of Lipid Research 46 (9): 1904-1913.

Li, Yan, Muhammad Farooq, Donglai Sheng, Chanchal Chandramouli, Tian Lan, Nilesh K Mahajan, R Manjunatha Kini, Yunhan Hong, Thomas Lisowsky, and Ruowen Ge. 2012. Augmenter of Liver Regeneration (alr) promotes liver outgrowth during zebrafish hepatogenesis. PLoS ONE 7 (1): e30835.

Lucero, Héctor a, and Phillips W Robbins. 2004. Lipid Rafts-protein Association and the Regulation of Protein Activity. Archives of Biochemistry and Biophysics 426 (2) (June 15): 208-24.

López-Casas, Pedro P, and Jesús Del Mazo. 2003. Regulation of Flotillin-1 in the Establishment of NIH-3T3 Cell-cell Interactions. FEBS Letters 555 (2): 223-228.

Lorent, Kristin, Sang-Yeob Yeo, Takaya Oda, Settara Chandrasekharappa, Ajay Chitnis, Randolph P Matthews, and Michael Pack. 2004. Inhibition of Jagged-mediated Notch Signaling Disrupts Zebrafish Biliary Development and Generates Multi-organ Defects Compatible with an Alagille Syndrome Phenocopy. Development Cambridge England 131 (22): 5753–5766.

Lv, Caixia, Travis J. Gould, Joerg Bewersdorf, and David Zenisek. 2012. High-Resolution Optical Imaging of Zebrafish Larval Ribbon Synapse Protein RIBEYE, RIM2, and CaV 1.4 by Stimulation Emission Depletion Microscopy. Microscopy and Microanalysis. 18(4):745-52.

Macara, Ian G. 2004. Parsing the Polarity Code. Nature Reviews. Molecular Cell Biology 5 (3) (March): 220–31.

Machta, Benjamin B., Stefanos Papanikolaou, James P. Sethna, and Sarah L. Veatch. 2011. Minimal Model of Plasma Membrane Heterogeneity Requires Coupling Cortical Actin to Criticality. Biophysical Journal 100 (7): 1668–1677.

193

Magenau, Astrid, Carola Benzing, Nicholas Proschogo, Anthony S Don, Leila Hejazi, Denuja Karunakaran, Wendy Jessup, and Katharina Gaus. 2011. Phagocytosis of IgG-Coated Polystyrene Beads by Macrophages Induces and Requires High Membrane Order. Traffic 12 (12): 1730–43.

Málaga-Trillo, Edward, Ute Laessing, Dirk M Lang, Axel Meyer, and Claudia A O Stuermer. 2002. Evolution of Duplicated Reggie Genes in Zebrafish and Goldfish. Journal of Molecular Evolution 54 (2): 235–245.

Mantena, S K, A L King, K K Andringa, H B Eccleston, and S M Bailey. 2008. Mitochondrial Dysfunction and Oxidative Stress in the Pathogenesis of Alcohol- and Obesity-induced Fatty Liver Diseases. Free Radical Biology and Medicine. 44 (7):1259-1272.

Martin-Belmonte, Fernando, Ama Gassama, Anirban Datta, Wei Yu, Ursula Rescher, Volker Gerke, and Keith Mostov. 2007. PTEN-mediated Apical Segregation of Phosphoinositides Controls Epithelial Morphogenesis Through Cdc42. Cell 128 (2): 383–385.

Martin J.Janiak, Donald M. Small, and G. Graham Shipley. 1979. Temperature and Compositional Dependence of the Structure of Hydrated Dimyristoyl Lecithin. Journal of Biological Chemistry. 254 (13): 6068-6078.

Masedunskas, Andrius, Natalie Porat-Shliom, Kamil Rechache, Myo-Pale’ Aye, and Roberto Weigert. 2012. Intravital Microscopy Reveals Differences in the Kinetics of Endocytic Pathways between Cell Cultures and Live Animals. Cells 1 (4): 1121–32.

Massey JB, Pownall HJ. 2005. The polar nature of 7-ketocholesterol determines its location within membrane domains and the kinetics of membrane microsolubilization by apolipoprotein A-I. Biochemistry 44: 10423–10433.

McCaffrey, L. M., and Macara, I. G. 2009. Widely conserved signaling pathways in the establishment of cell polarity. Cold Spring Harbor perspectives in biology, 1(2), 17.

McConnell, Harden M, and Marija Vrljic. 2003. Liquid-liquid Immiscibility in Membranes. Annual Review of Biophysics and Biomolecular Structure 32 (1): 469-492.

Meer, G V. 1989. Lipid Traffic in Animal Cells. Annual Review of Cell Biology 5: 247–275.

Mellman, Ira, and W James Nelson. 2008. Coordinated Protein Sorting, Targeting and Distribution in Polarized Cells. Nature Reviews Molecular Cell Biology 9 (11): 833–845.

Mentis MJ, Delalot D. 2005. Depression in Parkinson’s Disease. Adv Neurol 96: 26–41.

Milhiet, Pierre Emmanuel, Marie-Cécile Giocondi, and Christian Le Grimellec. 2002. Cholesterol Is Not Crucial for the Existence of Microdomains in Kidney Brush-border Membrane Models. The Journal of Biological Chemistry 277 (2) (January): 875-8.

M. J. Berridge and J. L. Oschman. 1972. Transporting Epithelia. Academic Press: 91–108.

194

Mondal, Mousumi, Bruno Mesmin, Sushmita Mukherjee, and Frederick R Maxfield. 2009. Sterols Are Mainly in the Cytoplasmic Leaflet of the Plasma Membrane and the Endocytic Recycling Compartment in CHO Cells. Molecular Biology of the Cell 20 (2): 581–588.

Morcos, Paul A. 2007. Achieving Targeted and Quantifiable Alteration of mRNA Splicing with Morpholino Oligos. Biochemical and Biophysical Research Communications 358 (2): 521–527.

Mostov, Keith E, and Marcel Verges and Yoram Altschuler. 2000. Membrane Traffic in Polarized Epithelial Cells. Current Opinion in Cell Biology 12 (Figure 1): 483–490.

Mostov, Keith, Tao Su, and Martin Ter Beest. 2003. Polarized Epithelial Membrane Traffic: Conservation and Plasticity. Nature Cell Biology 5 (4): 287–293.

Mouritsen, O G, and K Jørgensen. 1994. Dynamical Order and Disorder in Lipid Bilayers. Chemistry and Physics of Lipids 73 (1-2): 3–25.

Mukherjee, S, and F R Maxfield. 2000. Role of Membrane Organization and Membrane Domains in Endocytic Lipid Trafficking. Traffic Copenhagen Denmark 1 (3): 203–211.

Mueller, Veronika, Alf Honigmann, Christian Ringemann, Rebecca Medda, G??nter Schwarzmann, and Christian Eggeling. 2013. FCS in STED Microscopy: Studying the Nanoscale of Lipid Membrane Dynamics. Methods in Enzymology 519: 1–38.

Munderloh, Christina, Gonzalo P Solis, Vsevolod Bodrikov, Friederike A Jaeger, Marianne Wiechers, Edward Málaga-Trillo, and Claudia A O Stuermer. 2009. Reggies/flotillins Regulate Retinal Axon Regeneration in the Zebrafish Optic Nerve and Differentiation of Hippocampal and N2a Neurons. The Journal of Neuroscience 29 (20): 6607–6615.

Munro, Sean. 2003. Lipid Rafts : Elusive or Illusive ?. Cell 115: 377–388.

Murata, M, J Peränen, R Schreiner, F Wieland, T V Kurzchalia, and K Simons. 1995. VIP21/caveolin Is a Cholesterol-Binding Protein. Proceedings of the National Academy of Sciences of the United States of America 92 (22): 10339–10343.

Nasevicius, A, and S C Ekker. 2000. Effective Targeted Gene ‘Knockdown’ in Zebrafish. Nature Genetics 26 (2): 216–220.

Nelson, W James. 2009. Remodeling Epithelial Cell Organization: Transitions Between Front– Rear and Apical–Basal Polarity. Cold Spring Harbor Perspectives in Biology 1 (1): a000513.

Ng, Annie N Y, Tanya A De Jong-Curtain, David J Mawdsley, Sara J White, Jimann Shin, Bruce Appel, P Duc Si Dong, Didier Y R Stainier, and Joan K Heath. 2005. Formation of the Digestive System in Zebrafish: III. Intestinal Epithelium Morphogenesis. Developmental Biology 286 (1): 114–135.

Nichols, B. J., and G. P. Otto. 2011. The Roles of Flotillin Microdomains - Endocytosis and Beyond. Journal of Cell Science. 124, 3933-3940.

195

Nüsslein-Volhard, C. and Dahm, R. 2002. Zebrafish: a Practical Approach. New York: Oxford University Press.

Ober, Elke A, Holly A Field, and Didier Y R Stainier. 2003. From Endoderm Formation to Liver and Pancreas Development in Zebrafish. Mechanisms of Development 120 (1): 5–18.

Oehlers, Stefan H, Maria Vega Flores, Kazuhide S Okuda, Chris J Hall, Kathryn E Crosier, and Philip S Crosier. 2011. A Chemical Enterocolitis Model in Zebrafish Larvae That Is Dependent on Microbiota and Responsive to Pharmacological Agents. Developmental Dynamics an Official Publication of the American Association of Anatomists 240 (1): 288– 298.

Ohtani, Y, T Irie, K Uekama, K Fukunaga, and J Pitha. 1989. Differential Effects of Alpha-, Beta- and Gamma-Cyclodextrins on Human Erythrocytes. European Journal of Biochemistry / FEBS 186 (1-2): 17–22.

Ohvo, H, and J P Slotte. 1996. Cyclodextrin-Mediated Removal of Sterols from Monolayers: Effects of Sterol Structure and Phospholipids on Desorption Rate. Biochemistry 35 (24): 8018–8024.

Omori, Yoshihiro, and Jarema Malicki. 2006. Oko Meduzy and Related Crumbs Genes Are Determinants of Apical Cell Features in the Vertebrate Embryo. Current Biology 16 (10): 945–957.

Orlando, Kelly, and Wei Guo. 2009. Membrane Organization and Dynamics in Cell Polarity. Cold Spring Harbor Perspectives in Biology 1 (5) (November): a001321.

Ostasov, Pavel, Lenka Bourova, Lucie Hejnova, Jiri Novotny, and Petr Svoboda. 2007. Disruption of the Plasma Membrane Integrity by Cholesterol Depletion Impairs Effectiveness of TRH Receptor-Mediated Signal Transduction via G(q)/G(11)alpha Proteins. Journal of Receptor and Signal Transduction Research 27 (5-6): 335–352.

Owen, Dylan M., Astrid Magenau, Arindam Majumdar, and Katharina Gaus. 2010. Imaging Membrane Lipid Order in Whole, Living Vertebrate Organisms. Biophysical Journal 99 (1) (July): L7-L9.

Owen J S, Bruckdorfer K R, Day R C and McIntyre N. 1982. Decreased erythrocyte membrane fluidity and altered lipid composition in human liver disease. The Journal of Lipid Research, 23, 124-132.

Owen, Dylan M, Carles Rentero, Astrid Magenau, Ahmed Abu-Siniyeh, and Katharina Gaus. 2012. Quantitative Imaging of Membrane Lipid Order in Cells and Organisms. Nature Protocols 7 (1) (January): 24–35.

Owen, Dylan M, David J Williamson, Astrid Magenau, and Katharina Gaus. 2012. Sub- Resolution Lipid Domains Exist in the Plasma Membrane and Regulate Protein Diffusion and Distribution. Nature Communications 3: 1256.

196

Owen, Dylan M, Peter M P Lanigan, Christopher Dunsby, Ian Munro, David Grant, Mark a a Neil, Paul M W French, and Anthony I Magee. 2006. Fluorescence Lifetime Imaging Provides Enhanced Contrast When Imaging the Phase-sensitive Dye di-4-ANEPPDHQ in Model Membranes and Live Cells. Biophysical Journal 90 (11) (June): L80-2.

Owen, Dylan M, Stephane Oddos, Sunil Kumar, Daniel M Davis, Mark A A Neil, Paul M W French, Michael L Dustin, Anthony I Magee, and Marek Cebecauer. 2010. High Plasma Membrane Lipid Order Imaged at the Immunological Synapse Periphery in Live T Cells. Molecular Membrane Biology 27 (4-6): 178–189.

Pack, M, L Solnica-Krezel, J Malicki, S C Neuhauss, A F Schier, D L Stemple, W Driever, and M C Fishman. 1996. Mutations Affecting Development of Zebrafish Digestive Organs. Development Cambridge England 123: 321–328.

Palazzo, Alexander F, Christina H Eng, David D Schlaepfer, Eugene E Marcantonio, and Gregg G Gundersen. 2004. Localized Stabilization of Microtubules by Integrin- and FAK- facilitated Rho Signaling. Science 303 (5659): 836-839.

Parasassi, T, A M Giusti, M Raimondi, and E Gratton. 1995. Abrupt Modifications of Phospholipid Bilayer Properties at Critical Cholesterol Concentrations. Biophysical Journal 68 (5): 1895-1902.

Parasassi, T, E Gratton, W M Yu, P Wilson, and M Levi. 1997. Two-photon Fluorescence Microscopy of Laurdan Generalized Polarization Domains in Model and Natural Membranes. Biophysical Journal 72 (6) (June): 2413-29.

Parasassi, T, G De Stasio, A D'Ubaldo, and E Gratton. 1990. Phase Fluctuation in Phospholipid Membranes Revealed by Laurdan Fluorescence. Biophysical Journal 57 (6): 1179-1186.

Parasassi, Tiziana, and E K Krasnowska. 1998. Laurdan and Prodan as Polarity-sensitive Fluorescent Membrane Probes. Journal of Fluorescence 8 (4): 365–373.

Partridge, M, A Vincent, P Matthews, J Puma, D Stein, and J Summerton. 1996. A Simple Method for Delivering Morpholino Antisense Oligos into the Cytoplasm of Cells. Antisense & Nucleic Acid Drug Development 6 (3): 169–175.

Pearse, B M, and M S Robinson. 1990. Clathrin, Adaptors, and Sorting. Annual Review of Cell Biology 6 (January): 151–71.

Peer, Wendy Ann. 2011. The Plant Plasma Membrane. 19th ed. Monographs, Plant Cell.

Petronczki, M, and J A Knoblich. 2001. DmPAR-6 Directs Epithelial Polarity and Asymmetric Cell Division of Neuroblasts in Drosophila. Nature Cell Biology 3 (1): 43–49.

Pieczynski, Jay, and Ben Margolis. 2011. Protein Complexes That Control Renal Epithelial Polarity. American Journal of Physiology. Renal Physiology 300 (3): F589–F601.

Pike, L J, and J M Miller. 1998. Cholesterol Depletion Delocalizes Phosphatidylinositol Bisphosphate and Inhibits Hormone-stimulated Phosphatidylinositol Turnover. The Journal of Biological Chemistry 273 (35): 22298-22304. 197

Pike, Linda J. 2004. Lipid Rafts: Heterogeneity on the High Seas. The Biochemical Journal 378 (Pt 2) (March): 281-92.

Pike, Linda J. 2006. Rafts Defined: a Report on the Keystone Symposium on Lipid Rafts and Cell Function. Journal of Lipid Research 47 (7) (July): 1597-8.

Pike, Linda J. 2009. The Challenge of Lipid Rafts. Journal of Lipid Research 50 Suppl (April): S323-8.

Pike, Linda J, Xianlin Han, Koong-Nah Chung, and Richard W Gross. 2002. Lipid Rafts Are Enriched in Arachidonic Acid and Plasmenylethanolamine and Their Composition Is Independent of Caveolin-1 Expression: a Quantitative Electrospray Ionization/mass Spectrometric Analysis. Biochemistry 41 (6): 2075-2088.

Pitha, J, T Irie, P B Sklar, and J S Nye. 1988. Drug Solubilizers to Aid Pharmacologists: Amorphous Cyclodextrin Derivatives. Life Sciences 43 (6): 493–502.

Postlethwait, John, Angel Amores, William Cresko, Amy Singer, and Yi-Lin Yan. 2004. Subfunction Partitioning, the Teleost Radiation and the Annotation of the Human Genome. Trends in Genetics 20 (10): 481–490.

Pralle, A, P Keller, E L Florin, K Simons, and J K Hörber. 2000. Sphingolipid-Cholesterol Rafts Diffuse as Small Entities in the Plasma Membrane of Mammalian Cells. The Journal of Cell Biology 148 (5): 997–1008.

Praskova E, Voslarova E, Siroka Z, Macova S, Plhalova L, Bedanova I, Marsalek P, Pistekova V, Svobodova Z. 2011. Comparison of Acute Toxicity of Ketoprofen to Juvenile and Embryonic Stages of Danio Rerio. Neuro Endocrinol Lett 32 (1): 117–20. Progatzky, Fränze, Margaret J Dallman, and Cristina Lo Celso. 2013. From Seeing to Believing: Labelling Strategies for in Vivo Cell-Tracking Experiments. Interface Focus 3 (3): 20130001.

Prydz, K, and K Simons. 2001. Cholesterol Depletion Reduces Apical Transport Capacity in Epithelial Madin-Darby Canine Kidney Cells. The Biochemical Journal 357 (Pt 1): 11–15.

Quinn, Ronald J., David G. Le Couteur, Victoria C. Cogger, Bård Smedsrød, Gregory P. McNerney, Dmitri Svistounov, Dylan M. Owen, et al. 2012. The relationship between fenestrations, sieve plates and rafts in liver sinusoidal endothelial cells. PLoS ONE. 7(9): e46134.

Rajendran, Lawrence, Julia Beckmann, Astrid Magenau, Eva-Maria Boneberg, Katharina Gaus, Antonella Viola, Bernd Giebel, and Harald Illges. 2009. Flotillins are involved in the polarization of primitive and mature hematopoietic cells. PloS One 4 (12): e8290.

Rajendran, Lawrence, Madhan Masilamani, Samuel Solomon, Ritva Tikkanen, Claudia A O Stuermer, Helmut Plattner, and Harald Illges. 2003. Asymmetric Localization of Flotillins/Reggies in Preassembled Platforms Confers Inherent Polarity to Hematopoietic Cells. Proceedings of the National Academy of Sciences of the United States of America 100 (14): 8241–8246.

198

Ramstedt, Bodil, and J Peter Slotte. 2002. Membrane Properties of Sphingomyelins. FEBS Letters 531 (1): 33-37.

Ramstedt, Bodil, and J Peter Slotte. 2006. Sphingolipids and the Formation of Sterol-enriched Ordered Membrane Domains. Biochimica Et Biophysica Acta 1758 (12): 1945-1956.

Rankin, Brian R., Gael Moneron, Christian A. Wurm, Jessica C. Nelson, Arne Walter, Dirk Schwarzer, JÖrg Schroeder, Daniel A. ColÓn-Ramos, and Stefan W. Hell. 2011. Nanoscopy in a Living Multicellular Organism Expressing GFP. Biophysical Journal 100 (12).

Rentero, C, T Zech, C M Quinn, K Engelhardt, D Williamson, T Grewal, W Jessup, T Harder, and K Gaus. 2008. Functional Implications of Plasma Membrane Condensation for T Cell Activation. Plos One 3 (5).

Reimschuessel R., Ferguson H. W. 2006. Kidney. In Systemic Pathology of Fish, (2) 91–118.

Richard IH, Schiffer RB, Kurlan R. 2005. Anxiety and Parkinson's disease. Disorders in Parkinson's disease. Adv Neurol. 96:42-55.

Rinia, H A, M M Snel, J P Van Der Eerden, and B De Kruijff. 2001. Visualizing Detergent Resistant Domains in Model Membranes with Atomic Force Microscopy. FEBS Letters 501 (1): 92–96.

Rodriguez-boulan, Enrique, and W James Nelson. 1989. Morphogenesis of the Polarized Epithelial Cell Phenotype. 245 (4919): 718–725.

Roduit, Charles, F Gisou van der Goot, Paolo De Los Rios, Alexandre Yersin, Pascal Steiner, Giovanni Dietler, Stefan Catsicas, Frank Lafont, and Sandor Kasas. 2008. Elastic Membrane Heterogeneity of Living Cells Revealed by Stiff Nanoscale Membrane Domains. Biophysical Journal 94 (4): 1521–1532.

Rosen, Jonathan N, Michael F Sweeney, and John D Mably. 2009. Microinjection of Zebrafish Embryos to Analyze Gene Function. Journal of Visualized Experiments 127 (25): 2–5.

Rothman, James E. 1994. Mechanisms of Intracellular Protein Transport. Nature: 55–63.

Sadava D, Hillis DM, Heller HC, Berenbaum MR. 2011. Life: The Science of Biology. 9 th. San Francisco: Freeman.

Sako, Y, S Minoghchi, and T Yanagida. 2000. Single-Molecule Imaging of EGFR Signalling on the Surface of Living Cells. Nature Cell Biology 2 (3): 168–172.

Saltiel, A R, and C R Kahn. 2001. Insulin Signalling and the Regulation of Glucose and Lipid Metabolism. Nature 414 (6865): 799–806.

Samuel, B U, N Mohandas, T Harrison, H McManus, W Rosse, M Reid, and K Haldar. 2001. The Role of Cholesterol and Glycosylphosphatidylinositol-Anchored Proteins of Erythrocyte Rafts in Regulating Raft Protein Content and Malarial Infection. The Journal of Biological Chemistry 276 (31): 29319–29329. 199

Saslowsky, David E, Jin Ah Cho, Himani Chinnapen, Ramiro H Massol, Daniel J-F Chinnapen, Jessica S Wagner, Heidi E De Luca, Wendy Kam, Barry H Paw, and Wayne I Lencer. 2010. Intoxication of Zebrafish and Mammalian Cells by Cholera Toxin Depends on the Flotillin/reggie Proteins but Not Derlin-1 or -2. The Journal of Clinical Investigation 120 (12): 4399–4409.

Saxton, M J, and K Jacobson. 1997. Single-Particle Tracking: Applications to Membrane Dynamics. Annual Review of Biophysics and Biomolecular Structure 26: 373–399.

Scheiffele, P, A Rietveld, T Wilk, and K Simons. 1999. Influenza Viruses Select Ordered Lipid Domains During Budding from the Plasma Membrane. The Journal of Biological Chemistry 274 (4): 2038-2044.

Schlegel, Amnon. 2012. Studying Non-alcoholic Fatty Liver Disease with Zebrafish: a Confluence of Optics, Genetics, and Physiology. Cellular and Molecular Life Sciences. 69 (23): 3953-3961.

Schlüter, Marc A, Catherine S Pfarr, Jay Pieczynski, Eileen L Whiteman, Toby W Hurd, Shuling Fan, Chia-Jen Liu, and Ben Margolis. 2009. Trafficking of Crumbs3 During Cytokinesis Is Crucial for Lumen Formation. Molecular Biology of the Cell 20 (22): 4652– 4663.

Schmidt, T, and GJ Schütz. 1995. Characterization of Photophysics and Mobility of Single Molecules in a Fluid Lipid Membrane. The Journal of Physical Chemistry 99 (49): 17662– 17668.

Schuck, Sebastian, Masanori Honsho, Kim Ekroos, Andrej Shevchenko, and Kai Simons. 2003. Resistance of Cell Membranes to Different Detergents. Proceedings of the National Academy of Sciences of the United States of America 100 (10): 5795-5800.

Selkoe, Dennis J. 2001. Alzheimer’s Disease : Genes, Proteins, and Therapy. Physiol Rev 81 (2): 741–766.

Sengupta, Prabuddha, Adam Hammond, David Holowka, and Barbara Baird. 2009. Structural Determinants for Partitioning of Lipids and Proteins Between Coexisting Fluid Phases in Giant Plasma Membrane Vesicles. Biochim Biophys Acta. 1778 (1): 20-32.

Seth, Asha, Derek L Stemple, and Inês Barroso. 2013. The Emerging Use of Zebrafish to Model Metabolic Disease. Disease Models & Mechanisms 6 (5): 1080–8.

Sharma, Pranav, Rajat Varma, R.C Sarasij, Karine Gousset, G Krishnamoorthy, Madan Rao, and Satyajit Mayor. 2004. Nanoscale Organization of Multiple GPI-Anchored Proteins in Living Cell Membranes. Cell 116 (4) (February): 577-589.

Shashkin, Pavel, Bojan Dragulev, and Klaus Ley. 2005. Macrophage Differentiation to Foam Cells. Current Pharmaceutical Design 11 (23): 3061-3072.

Shim, Katherine. 2011. Vibratome Sectioning for Enhanced Preservation of the Cytoarchitecture of the Mammalian Organ of Corti. Journal of Visualized Experiments JoVE (52): 2–5. 200

Shin, Kunyoo, Vanessa C Fogg, and Ben Margolis. 2006. Tight Junctions and Cell Polarity. Annual Review of Cell and Developmental Biology 22 (June): 207–235.

Shogomori, Hidehiko, and Deborah A Brown. 2003. Use of Detergents to Study Membrane Rafts: The Good, the Bad, and the Ugly. Biological Chemistry 384 (9): 1259-1263.

Schroeder, R J, S N Ahmed, Y Zhu, E London, and D A Brown. 1998. Cholesterol and Sphingolipid Enhance the Triton X-100 Insolubility of Glycosylphosphatidylinositol- Anchored Proteins by Promoting the Formation of Detergent-Insoluble Ordered Membrane Domains. The Journal of Biological Chemistry 273 (2): 1150–1157.

Shi, Xiaoli, Xuejie Zhang, Tie Xia, and Xiaohong Fang. 2012. Living Cell Study at the Single- Molecule and Single-Cell Levels by Atomic Force Microscopy. Nanomedicine (London, England) 7 (10): 1625–37.

Shivas, Jessica M., Holly A. Morrison, David Bilder, and Ahna R. Skop. 2010. Polarity and Endocytosis: Reciprocal Regulation. Trends in Cell Biology. 20(8):445-52.

Silvius, John R. 2003. Role of Cholesterol in Lipid Raft Formation: Lessons from Lipid Model Systems. Biochimica et Biophysica Acta 1610 (2): 174–183.

Simons, K, and Stephen D. Fuller. Cell surface polarity in epithelia. Annual review of cell biology 1, no. 1 (1985): 243-288.

Simons, K, and E Ikonen. 1997. Functional Rafts in Cell Membranes. Nature 387 (6633): 569- 572.

Simons, K, and D Toomre. 2000. Lipid Rafts and Signal Transduction. Nature Reviews. Molecular Cell Biology 1 (1) (October): 31-9.

Simons, Kai, and Gerrit Van Meers. 1988. Perspectives in Biochemistry Lipid Sorting in Epithelial Cells. Biochemistry 27 (17): 6197-6202.

Simons, Kai, and Winchil L C Vaz. 2004. Model Systems, Lipid Rafts, and Cell Membranes. Annual Review of Biophysics and Biomolecular Structure 33 (January): 269-95.

Slaughter, Ndaisha, Isett Laux, Xiaolin Tu, Julian Whitelegge, Xiaoming Zhu, Rita Effros, Perry Bickel, and Andre Nel. 2003. The Flotillins Are Integral Membrane Proteins in Lipid Rafts That Contain TCR-Associated Signaling Components: Implications for T-Cell Activation. Clinical Immunology (Orlando, Fla.) 108 (2): 138–151.

Smart, Eric J, Robert A De Rose, and Steven A Farber. 2004. Annexin 2-Caveolin 1 Complex Is a Target of Ezetimibe and Regulates Intestinal Cholesterol Transport. Proceedings of the National Academy of Sciences of the United States of America 101 (10): 3450–3455.

Smith, A E, and A Helenius. 2004. How Viruses Enter Animal Cells. Science 304 (5668): 237- 242.

201

Solis, Gonzalo P, Maja Hoegg, Christina Munderloh, Yvonne Schrock, Edward Malaga-Trillo, Eric Rivera-Milla, and Claudia A O Stuermer. 2007. Reggie/flotillin Proteins Are Organized into Stable Tetramers in Membrane Microdomains. The Biochemical Journal 403 (2): 313-322.

Song, Libing, Hui Gong, Chuyong Lin, Chanjuan Wang, Liping Liu, Jueheng Wu, Mengfeng Li, and Jun Li. 2012. Flotillin-1 Promotes Tumor Necrosis Factor-Α Receptor Signaling and Activation of NF-Κb in Esophageal Squamous Cell Carcinoma Cells. Gastroenterology 143 (4).

Sot, Jesús, M Isabel Collado, José L R Arrondo, Alicia Alonso, and Félix M Goñi. 2002. Triton X-100-Resistant Bilayers: Effect of Lipid Composition and Relevance to the Raft Phenomenon. Langmuir 18 (7): 2828-2835.

Sotillos, Sol, María Teresa Díaz-Meco, Eva Caminero, Jorge Moscat, and Sonsoles Campuzano. 2004. DaPKC-Dependent Phosphorylation of Crumbs Is Required for Epithelial Cell Polarity in Drosophila. The Journal of Cell Biology 166 (4): 549–557.

Spence, Rowena, Gabriele Gerlach, Christian Lawrence, and Carl Smith. 2008. The Behaviour and Ecology of the Zebrafish, Danio Rerio. Biological Reviews of the Cambridge Philosophical Society 83 (1): 13-34.

Stainier, D Y. 2001. Zebrafish Genetics and Vertebrate Heart Formation. Nature Reviews Genetics 2 (1): 39–48.

Steck, Theodore L, Jin Ye, and Yvonne Lange. 2002. Probing Red Cell Membrane Cholesterol Movement with Cyclodextrin. Biophysical Journal 83 (4): 2118–2125.

St Johnston, Daniel, and Julie Ahringer. 2010. Cell Polarity in Eggs and Epithelia: Parallels and Diversity. Cell 141 (5): 757–774.

Stoletov, Konstantin, Longhou Fang, Soo-Ho Choi, Karsten Hartvigsen, Lotte F Hansen, Chris Hall, Jennifer Pattison, et al. 2009. Vascular Lipid Accumulation, Lipoprotein Oxidation, and Macrophage Lipid Uptake in Hypercholesterolemic Zebrafish. Circulation Research 104 (8) (April): 952-60.

Strazzabosco, Mario, and Stefan Somlo. 2011. Polycystic Liver Diseases: Congenital Disorders of Cholangiocyte Signaling. Gastroenterology 140 (7): 1855–1859, 1859.e1.

Stuermer, C. A. O. 2011. Reggie/flotillin and the targeted delivery of cargo. Journal of Neurochemistry, 116: 708–713.

Summerton, J. 1999. Morpholino Antisense Oligomers: The Case for an RNase H-Independent Structural Type. Biochimica et Biophysica Acta 1489 (1): 141–158.

Summerton, J, D Stein, S B Huang, P Matthews, D Weller, and M Partridge. 1997. Morpholino and Phosphorothioate Antisense Oligomers Compared in Cell-Free and in-Cell Systems. Antisense & Nucleic Acid Drug Development 7 (2): 63–70.

202

Summerton, J, and D Weller. 1997. Morpholino Antisense Oligomers: Design, Preparation, and Properties. Antisense & Nucleic Acid Drug Development 7 (3): 187–195.

Sun, Zhaoxia, Adam Amsterdam, Gregory J Pazour, Douglas G Cole, Mark S Miller, and Nancy Hopkins. 2004. A Genetic Screen in Zebrafish Identifies Cilia Genes as a Principal Cause of Cystic Kidney. Development Cambridge England 131 (16): 4085–4093.

Surma, Michal A., Christian Klose, and Kai Simons. 2012. Lipid-Dependent Protein Sorting at the Trans-Golgi Network. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids. 1821(8):1059-67.

Suzuki, Atsushi, and Shigeo Ohno. 2006. The PAR-aPKC System: Lessons in Polarity. Journal of Cell Science 119 (Pt 6) (March 15): 979–87.

Suzuki, Atsushi, Tomoyuki Yamanaka, Tomonori Hirose, Naoyuki Manabe, Keiko Mizuno, Miki Shimizu, Kazunori Akimoto, Yasushi Izumi, Tetsuo Ohnishi, and Shigeo Ohno. 2001. Atypical Protein Kinase C Is Involved in the Evolutionarily Conserved Par Protein Complex and Plays a Critical Role in Establishing Epithelia-Specific Junctional Structures. The Journal of Cell Biology 152 (6): 1183–1196.

Tabuse, Y, Y Izumi, F Piano, K J Kemphues, J Miwa, and S Ohno. 1998. Atypical Protein Kinase C Cooperates with PAR-3 to Establish Embryonic Polarity in Caenorhabditis Elegans. Development Cambridge England 125 (18): 3607–3614.

Takahashi, Eriko, Osamu Inanami, Toshio Ohta, Akira Matsuda, and Mikinori Kuwabara. 2006. Lipid Raft Disruption Prevents Apoptosis Induced by 2-chloro-2'-deoxyadenosine (Cladribine) in Leukemia Cell Lines. Leukemia Research 30 (12): 1555-1561.

Takeuchi, M, H Miyamoto, Y Sako, H Komizu, and A Kusumi. 1998. Structure of the Erythrocyte Membrane Skeleton as Observed by Atomic Force Microscopy. Biophysical Journal 74 (5): 2171-2183.

Tanentzapf, Guy, and Ulrich Tepass. 2003. Interactions between the Crumbs, Lethal Giant Larvae and Bazooka Pathways in Epithelial Polarization. Nature Cell Biology 5 (1): 46– 52.

Tao, Ting, and Jinrong Peng. 2009. Liver Development in Zebrafish (Danio Rerio). Journal of Genetics and Genomics = Yi Chuan Xue Bao 36 (6): 325–334.

Tepass, Ulrich. 2012. The Apical Polarity Protein Network in Drosophila Epithelial Cells: Regulation of Polarity, Junctions, Morphogenesis, Cell Growth, and Survival. Annual Review of Cell and Developmental Biology 28: 655–85.

Thiele, C, and W B Huttner. 1998. Protein and Lipid Sorting from the trans-Golgi Network to Secretory Granules-recent Developments. Seminars in Cell Developmental Biology 9 (5): 511–516.

Thisse, Christine, and Leonard I Zon. 2002. Organogenesis--heart and Blood Formation from the Zebrafish Point of View. Science 295 (5554): 457–462.

203

Tietz Bogert, Pamela S, Bing Q Huang, Sergio A Gradilone, Tetyana V Masyuk, Gary L Moulder, Stephen C Ekker, and Nicholas F Larusso. 2013. The Zebrafish as a Model to Study Polycystic Liver Disease. Zebrafish 10 (00): 1–7.

Traian D, M. M., F. L. G. Flecha, and V. Levi. 2012. Imaging Lipid Lateral Organization in Membranes with C-Laurdan in a Confocal Microscope. The Journal of Lipid Research.

Van Der Goot, F G, and T Harder. 2001. Raft Membrane Domains: From a Liquid-ordered Membrane Phase to a Site of Pathogen Attack. Seminars in Immunology 13 (2): 89-97.

Van Der Ven, Leo T M, Henrik Holbech, Martina Fenske, Evert-Jan Van Den Brandhof, Francijntje K Gielis-Proper, and Piet W Wester. 2003. Vitellogenin Expression in Zebrafish Danio Rerio: Evaluation by Histochemistry, Immunohistochemistry, and in Situ mRNA Hybridisation. Aquatic Toxicology Amsterdam Netherlands 65 (1): 1–11.

Van Meer, G. 1998. Lipids of the Golgi Membrane. Trends in Cell Biology 8 (1): 29-33.

Van Meer, G, and K Simons. 1982. Viruses Budding from Either the Apical or the Basolateral Plasma Membrane Domain of MDCK Cells Have Unique Phospholipid Compositions. The EMBO Journal 1 (7): 847–852.

Vascotto, S G, Y Beckham, and G M Kelly. 1997. The Zebrafish's Swim to Fame as an Experimental Model in Biology. Biochemistry and Cell Biology Biochimie Et Biologie Cellulaire 75 (5): 479-485.

Veatch, Sarah L, and Sarah L Keller. 2002. Organization in Lipid Membranes Containing Cholesterol. Physical Review Letters 89 (26): 268101.

Vicidomini, Giuseppe, Gael Moneron, Kyu Y Han, Volker Westphal, Haisen Ta, Matthias Reuss, Johann Engelhardt, Christian Eggeling, and Stefan W Hell. 2011. Sharper Low- Power STED Nanoscopy by Time Gating. Nature Methods 8 (7): 571–573.

Vieira, Otilia V, Katharina Gaus, Paul Verkade, Joachim Fullekrug, Winchil L C Vaz, and Kai Simons. 2006. FAPP2, Cilium Formation, and Compartmentalization of the Apical Membrane in Polarized Madin-Darby Canine Kidney (MDCK) Cells. Proceedings of the National Academy of Sciences of the United States of America 103 (49): 18556–18561.

Vihtelic TS, Doro CJ, Hyde DR. 1991. Cloning and characterization of six zebrafish photoreceptor opsin cDNAs and immunolocalization of their corresponding proteins. Vis Neurosci; 16(3):571-85.

Vishwajeet Puri, Rikio Watanabe, Michel Dominguez, Xiaofeng Sun, Christine L. Wheatley, David L. Marks & Richard E. Pagano. 1999. Cholesterol Modulates Membrane Traffic Along the Endocytic Pathway in Sphingolipid-storage Diseases. Nature Cell Biology 1.

Von Heijne G, Rees D. 2008. Membranes: reading between the lines. Current Opinion in Structural Biology, 18:403-405.

204

Von Philipsborn, Anne C, Anna Ferrer-Vaquer, Eric Rivera-Milla, Claudia A O Stuermer, and Edward Málaga-Trillo. 2005. Restricted Expression of Reggie Genes and Proteins During Early Zebrafish Development. Journal of Comparative Neurology 482 (3): 257-272.

Wallace, K N, S Akhter, E M Smith, K Lorent, and M Pack. 2005. Intestinal Growth and Differentiation in Zebrafish. Mechanisms of Development 122 (2): 157–173.

Walther, Rhian F, and Franck Pichaud. 2010. Crumbs/DaPKC-Dependent Apical Exclusion of Bazooka Promotes Photoreceptor Polarity Remodeling. Current Biology : CB 20 (12): 1065–1074.

Weber, G, and F.J. Farris. 1979. Synthesis and Spectral Properties of a Hydrophobic Fluorescent Probe: 6-Propionyl-2-(dimethylamino) Naphthalene. Biochemistry 18 (14): 3075–3078.

Wei, Xiangyun, Yan Cheng, Yiying Luo, Xiaohai Shi, Scott Nelson, and David R Hyde. 2004. The Zebrafish Pard3 Ortholog Is Required for Separation of the Eye Fields and Retinal Lamination. Developmental Biology 269 (1): 286–301.

Westerfield, M. 1995. The Zebrafish Book. Eugene: University of Oregon Press.

White, Richard Mark, Anna Sessa, Christopher Burke, Teresa Bowman, Jocelyn LeBlanc, Craig Ceol, Caitlin Bourque, et al. 2008. Transparent Adult Zebrafish as a Tool for in Vivo Transplantation Analysis. Cell Stem Cell 2 (2): 183–189.

Whittaker, J.R. 1966. An Analysis of Melanogenesis in Differentiating Pigment Cells of Ascidian Embryos. Developmental Biology: 14:1–39.

Willig, Katrin I, Benjamin Harke, Rebecca Medda, and Stefan W Hell. 2007. STED Microscopy with Continuous Wave Beams. Nature Methods 4 (11): 915–918.

Wingert, R A, and A J Davidson. 2008. The Zebrafish Pronephros: a Model to Study Nephron Segmentation. Kidney International 73 (10): 1120–1127.

Wingert, Rebecca A, Rori Selleck, Jing Yu, Huai-Dong Song, Zhu Chen, Anhua Song, Yi Zhou, et al. 2007. The Cdx Genes and Retinoic Acid Control the Positioning and Segmentation of the Zebrafish Pronephros. PLoS Genetics 3 (10): 1922–1938.

Xing, Yanghui, Yan Gu, Li-Chong Xu, Christopher A Siedlecki, Henry J Donahue, and Jun You. 2011. Effects of Membrane Cholesterol Depletion and GPI-Anchored Protein Reduction on Osteoblastic Mechanotransduction. Journal of Cellular Physiology 226 (9): 2350–2359.

Xingjie, Liang; Youguo, Huang. 2001. Alteration of membrane lipid biophysical properties and resistance of human lung adenocarcinoma A549 cells to cisplatin. Science in China Series C: Life Sciences 44: 25-32.

Yuan, Chunbo, Jennifer Furlong, Pierre Burgos, and Linda J Johnston. 2002. The Size of Lipid Rafts: An Atomic Force Microscopy Study of Ganglioside GM1 Domains in sphingomyelin/DOPC/cholesterol Membranes. Biophysical Journal 82 (5): 2526–2535. 205

Yuan, Shiaulou, and Zhaoxia Sun. 2009. Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos. Journal of Visualized Experiments 27 (27): 5–7.

Zeljka Korade and Anne K. Kenworthy. 2009. Lipid Rafts, Cholesterol, and the Brain. Molecular Physiology 55 (8): 1265-1273.

Zeng, Youchun, Nengbing Tao, Koong-Nah Chung, John E Heuser, and Douglas M Lublin. 2003. Endocytosis of Oxidized Low Density Lipoprotein Through Scavenger Receptor CD36 Utilizes a Lipid Raft Pathway That Does Not Require Caveolin-1. The Journal of Biological Chemistry 278 (46): 45931–45936.

Zhang, Jie, Andrew Pekosz, and Robert A Lamb. 2000. Influenza Virus Assembly and Lipid Raft Microdomains: a Role for the Cytoplasmic Tails of the Spike Glycoproteins. Journal of Virology 74 (10): 4634-4644.

Zhou, Weibin, Rudrick C Boucher, Frank Bollig, Christoph Englert, and Friedhelm Hildebrandt. 2010. Characterization of Mesonephric Development and Regeneration Using Transgenic Zebrafish. American Journal of Physiology. Renal Physiology 299 (5): F1040–F1047.

Zidovetzki, Raphael, and Irena Levitan. 2007. Use of Cyclodextrins to Manipulate Plasma Membrane Cholesterol Content: Evidence, Misconceptions and Control Strategies. Biochimica et Biophysica Acta 1768 (6): 1311–1324.

206

Appendix A

Publications arising from this study

Abu siniyeh. A, Owen .D, Majumdar. A, Gaus. K. Membrane order and distribution of polarity proteins are interconnected in polarized epithelial cells in the developing zebrafish larvae (not published yet).

207