Sphingolipid Biosynthesis, Metabolism and Actions in Placental Trophoblast Differentiation

Sphingolipid biosynthesis, metabolism and actions in placental trophoblast differentiation

Ambika Singh

BSc, MSc (Hons)

This thesis is presented in 2012 for the degree of

DOCTOR OF PHILOSOPHY (PhD) Of The University of Western Australia,

School of Women’s and Infants’ Health, Faculty of Medicine, Dentistry and Health Sciences & School of Anatomy and Human Biology, Faculty of Life and Physical Sciences

ABSTRACT

Formation and maintenance of the syncytiotrophoblast layer of the human placenta involves the fusion and terminal differentiation of trophoblast cells, the specialized cell type of the placenta. This process is unique and although it has been actively investigated over recent years, it remains incompletely understood. Sphingolipids such as sphingosine and ceramide are important endogenous cellular components which are controlled at the level of synthesis, metabolism and distribution, and act as regulators of numerous cellular functions including apoptosis and differentiation. The role of sphingolipids in trophoblast apoptosis, differentiation and fusion has not been elucidated in detail, apart from a study by Johnstone et al. indicating a negative role of sphingosine-1-phosphate (S1P) in this process. A previous work from our laboratory also implicated a role for ceramide in this process. The aim of this project, therefore, was to examine the production, metabolism and actions of key sphingolipids - ceramide, sphingosine and S1P - during spontaneous trophoblast differentiation in vitro to determine their role(s) in the differentiation process. Since earlier findings in a cell line model had shown an association between intracellular levels of ceramide and the lipid transporter ABCG2 (also known as breast cancer resistance protein, BCRP), I also investigated the putative link between ABCG2 function, sphingolipid levels and trophoblast differentiation.

I established a model of syncytialization, adapted from previously published methods, using human cytotrophoblasts extracted from term placentas syncytialized over 7 days in culture. Characterization of this model showed that patterns of expression/production differentiation markers – human chorionic gonadotrophin (hCG), alkaline phosphatase and GCM1 – varied during the differentiation process and that differentiation and cell fusion were independent processes. Lipids were extracted during the differentiation process and analysed by liquid chromatography- mass spectrometry. Intracellular C16 ceramide levels increased modestly after 3 days in culture then declined to basal levels, accompanied by changes in expression of ceramide synthesizing (sphingomyelinase, ceramide synthase) and degrading (ceramidase) enzymes. Ceramidase was present at particularly high levels in syncytialized trophoblasts; inhibition of ceramidase reduced the degree of cell fusion, suggesting

i that this enzyme may be involved in maintaining the syncytial phenotype. Exposure of trophoblasts for 72 h to short chain C8 ceramide or sphingomyelinase to increase cellular levels of ceramide enhanced secretion of the differentiation marker hCG without affecting fusion or cell viability.

In contrast to ceramide, significant declines in intracellular sphingosine levels and protein levels of sphingosine kinase 1 (SPHK1, the rate-limiting enzyme for S1P production) were observed during trophoblast differentiation and fusion. Secreted S1P levels dropped steeply while hCG secretion levels and levels of intraceullular ceramide were maximum, before rising back to basal levels with syncytialization. Intracellular S1P levels were undetectable. Treating cells with either exogenous sphingosine, S1P or a specific SPHK1 inhibitor for up to 72 h in culture significantly inhibited trophoblast differentiation (measured as reduced hCG production); effects on other biochemical and morphological markers of differentiation were absent or inconsistent.

Inhibition of the ceramide- and S1P-responsive pathways, namely c-Jun N-terminal kinase (JNK), protein phosphatase 2A (PP2A), p38MAPK, phosphatidylinositol 3- kinase (PI3K) and ERK1/2 did not abolish the effects of these bioactive mediators, and JNK phosphorylation was unresponsive to these compounds. However, novel roles of protein kinase B (PKB)/Akt, a growth and differentiation factor, and JNK in trophoblast differentiation and fusion were identified in this study as indicated by spontaneous changes in their phosphorylation state with syncytialisation. Ceramide and S1P significantly inhibited phosphorylation of Akt, suggesting that this may be a novel mechanism through which trophoblast differentiation may be regulated by sphingolipids.

Overall, my findings suggest that changes in ceramide biosynthesis and metabolism play a significant role in modulating the biochemical and morphological features of trophoblast differentiation. The data support the notion that at least some aspects of trophoblast differentiation and fusion can be dissociated as indicated by the differences in response between morphological and biochemical differentiation markers. My finding that ceramide is an intracellular regulator able to exert differential effects on functional trophoblast differentiation and syncytialization raises questions regarding the significance of ceramide synthesis and metabolism in abnormal pregnancies.

ii These studies did not confirm the expected association between sphingolipids (ceramide and sphingosine-1-phosphate) and ABCG2 during trophoblast differentiation and syncytialization, and hence do not support a role for ABCG2 in trophoblast differentiation in the human placenta, in contrast to findings previously reported using the BeWo choriocarcinoma cell line.

In conclusion, the principal findings of this thesis suggest that sphingolipid homeostasis is regulated during trophoblast differentiation and fusion and that its dysregulation could result in defects in placental formation and function. Disturbances in sphingolipid homeostasis could, therefore, have deleterious consequences for pregnancy outcome and fetal well being, as seen in pregnancy complications such as preeclampsia which are characterized by aberrant placental syncytialization.

iii ACKNOWLEDGMENTS

I am extremely grateful to my supervisor Professor Jeff Keelan, whose patience, kindness, and academic expertise, have been invaluable to me. His compassionate support and constant guidance helped my thesis through to completion. An excellent mentor and role model. Studying in his group provided a brilliant training platform, offering a varied range of opportunities and developmental skills.

I am heartily thankful to my co-supervisor Professor Arun Dharmarajan, whose encouragement, guidance and support from the onset to the final level enabled me to develop an understanding of the subject and opening several networking opportunities.

The School of Women’s and Infants’ Health provided remarkable study and lab facilities with a bonus exceptional team to work with. I would like to thank the funding support of the Women’s and Infants’ Research Foundation (WIRF) at King Edward Memorial Hospital, University of Western Australia, Perth, and the Liggins Institute, University of Auckland, New Zealand.

I would like to thank Sonia Alix, Eric Thorstensen and Thomas Stoll for carrying out the challenging sphingolipid mass spectrometry assays successfully with utmost proficiency.

The intellectual support and feedback provided by fellow PhD student Irving Aye, is truly appreciated. His generosity and friendship contributed significantly towards completion and submission of this thesis. Other colleagues I would like to thank for their guidance include Biju Balakrishna, Denis Evseenko and Vijay Pandey.

I am indebted to my friends, Disha Saxena, Parul Chauhan and Kirthana Prabhakaran, for providing constant encouragement and motivation.

Ronak Sampat, for making a sterling effort in proof reading my thesis (coming from a non- science background). I am eternally grateful for the continuous motivation in making me strive for excellence and give this thesis my best shot. Thanks for having an enormous amount of faith in me, the invaluable words of wisdom and referral to value-adding books.

I owe my deepest gratitude to my beloved parents and brother, Pranay Singh. This thesis would not be possible minus their constant source of support. My pillar of strength, joy and guiding light. Thank you very much for creating an environment in which following this path seemed so natural, and for having tremendous patience.

iv ACHIEVEMENTS DURING PhD

Manuscripts and abstracts arising from this thesis

1. AT Singh, A Dharmarajan, ILMH Aye, JA Keelan. Role of ceramide biosynthesis and metabolism in the regulation of trophoblast differentiation and syncytialisation. Molecular and Cellular Endocrinology. In revision.

2. AT Singh, A Dharmarajan, ILMH Aye, JA Keelan. The Sphingosine-S1P pathway regulates trophoblast differentiation and syncytialization. Reproductive BioMedicine Online. 24: 224– 234.

3. A Al-Khan, ILMH Aye, AT Singh, et al.: IFPA Meeting 2010 Workshops Report II: Placental pathology; Trophoblast invasion; Fetal sex; Parasites and the placenta; Decidua and embryonic or fetal loss; Trophoblast differentiation and syncytialization. Placenta. Placenta. 2011; 32 (2): S90-S99.

4. AT Singh and JA Keelan: Faculty of 1000 Biology, 22 Apr 2010 http://f1000biology.com/article/id/3001967/evaluation.

5. AT Singh and JA Keelan: Faculty of 1000 Biology, 21 Apr 2010 http://f1000biology.com/article/id/2982958/evaluation.

6. AT Singh, ILMH Aye, JA Keelan. Transport of lipids by ABC proteins: Interactions and implications for cellular toxicity, viability and function. Chemico-Biological Interactions. 2009; 180: 327-339. Awarded as one of the journal's top ten most downloaded articles of 2009.

7. AT Singh, A Dharmarajan, JA Keelan: A novel role of ceramide, sphingosine and sphingosine-1-phosphate in trophoblast differentiation, 2009. Placenta; Vol 30: abstract P01.04.

8. AT Singh, A Dharmarajan, JA Keelan. The Cer-S1P rheostat regulates trophoblast differentiation, 2009. Reproductive Sciences; Vol 16 (3) (suppl): abstract 955.

v 9. AT Singh, JW Paxton, JA Keelan. Studies of the role of ABC transporters and sphingolipids in trophoblast differentiation, 2007. Endocrine Journal 54 (suppl): 131, abstract 208.

Scholarships and Awards

1. Y.W. (Charlie) Loke Travel Award, IFPA meeting (2010)

2. ANZPRA New Investigator Travel Award, IFPA meeting (2009)

3. The Australian Postgraduate Award (APA)

4. University of Western Australia Top-Up Scholarship (UPA)

5. Women’s and Infants’ Research Foundation Top-Up Scholarship (2008)

6. Liggins Institute Postgraduate Scholarship (2006-2007)

7. First prize for best Oral presentation at Society of Reproductive Biology and Comparative Endocrinology (SRBCE) Meeting, January 2009, India.

8. UWA School of Pathology and Medicine Prize for Oral Presentation at the Australian Society for Medical Research Meeting (ASMR), June 2008, Australia.

9. ASMR Travel Grant (2008)

10. Society of Reproductive Biology (SRB) Travel Grant (2007)

Conference Presentations

1. AT Singh, A Dharmarajan, JA Keelan (Oral Presentation). International Federation of Placental Associations (IFPA), October 2010. Santiago, Chile.

2. AT Singh, A Dharmarajan, JA Keelan (Poster Presentation). International Federation of Placental Associations (IFPA), October 2009. Adelaide, Australia.

3. AT Singh, A Dharmarajan, JA Keelan (Oral Presentation). The Endocrine and Reproductive Biology Society of WA (ERBSWA), April 2009. Perth, Australia.

vi 4. AT Singh, A Dharmarajan, JA Keelan (Poster Presentation). Society of Gynaecologic Investigation (SGI), March 2009. Glasgow, Scotland.

5. AT Singh, A Dharmarajan, JA Keelan (Oral Presentation). Society of Reproductive Biology and Comparative Endocrinology (SRBCE), January 2009. Hyderabad, India.

6. AT Singh, JW Paxton, A Dharmarajan, JA Keelan (Oral Presentation). Australian Health and Medical Research Congress (AHMRC), November 2008. Brisbane, Australia.

7. AT Singh, JW Paxton, A Dharmarajan, JA Keelan (Oral Presentation). Postgrad Student Expo, July 2008. Perth, Australia.

8. AT Singh, JW Paxton, A Dharmarajan, JA Keelan (Oral Presentation). Australian Society for Medical Research (ASMR) Meeting, June 2008. Perth, Australia.

9. AT Singh, JW Paxton, JA Keelan (Oral Presentation). Exposure07, October 2007. Auckland, New Zealand.

10. AT Singh, JW Paxton, JA Keelan (Oral Presentation). HealtheX – the inaugural exposition Celebrating Student Research, September 2007. Auckland, New Zealand.

11. AT Singh, JW Paxton, JA Keelan (Oral Presentation). SRB Young Investigator nominee. 38th Annual Conference of the Society for Reproductive Biology, September 2007. Christchurch, New Zealand.

12. AT Singh, JA Keelan, F Sieg (Poster Presentation). ASCEPT NZ Meeting, December 2006. Auckland, New Zealand

vii Invited Speaker

1. AT Singh, A Dharmarajan, JA Keelan. ‘Trophoblast differentiation and syncytialization’ workshop. International Federation of Placental Associations (IFPA), October 2010. Santiago, Chile.

2. AT Singh, A Dharmarajan, JA Keelan. ‘Bioactive lipids in placenta’ workshop. International Federation of Placental Associations (IFPA), October 2009. Adelaide, Australia.

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CONTENTS

ABSTRACT ...... I

ACKNOWLEDGMENTS ...... IV

ACHIEVEMENTS DURING PHD ...... V

Manuscripts and Abstracts from this thesis ...... v

Scholarships and Awards ...... vi

Conference Presentations ...... vi

Invited Speaker ...... viii

CONTENTS ...... IX

LIST OF FIGURES ...... XIII

LIST OF TABLES ...... XVIII

ABBREVIATIONS ...... XIX

CHAPTER 1. INTRODUCTION ...... 1

1.1. Trophoblast differentiation and syncytial formation ...... 1 1.1.1. Hormones, growth factors and cytokines ...... 6 1.1.2. Membrane proteins ...... 9 1.1.3. Protein kinase ...... 13 1.1.4. Proteases ...... 13 1.1.5. Physicochemical factors ...... 15 1.1.6. Membrane architecture ...... 16 1.1.7. Transcription factors ...... 18 1.1.8 Summary ...... 19

1.2. Sphingolipids ...... 20 1.2.1. Ceramide ...... 22 i) Synthesis and Metabolism ...... 22 ii) Structural role ...... 26 iii) Ceramide as a second messenger ...... 28 1.2.2. Sphingosine (Sph) ...... 30 1.2.3. Sphingosine-1-phosphate (S1P) ...... 33 1.2.4. Regulation of ABC transporter function by SPLs ...... 39

1.3. ABC transporters ...... 41

1.3.1. Endogenous SPL substrates of ABC transporters ...... 42 i) Mechanism of lipid efflux ...... 42 ii) Efflux and trafficking of SPLs ...... 43 MDR1 ...... 43 MRP1 ...... 44 ABCA family ...... 44 1.3.2. Effects of membrane SPLs on ABC transporter activity ...... 46 i) MDR1 ...... 46 ii) ABCA1 ...... 47 iii) ABCG family ...... 47

1.4. ABC transporters, SPLs and trophoblast differentiation ...... 48

1.5. Summary, aims and hypotheses ...... 50

CHAPTER 2. MATERIALS AND METHODS ...... 53

2.1. Reagents ...... 53

2.2. Isolation and culture of term human placental trophoblast cells ...... 57 2.2.1. Buffers and Solutions...... 57 2.2.2. Trophoblast isolation ...... 57

2.3. Gene Expression Analyses ...... 60 2.3.1. RNA extraction ...... 60 2.3.2. Genomic DNA removal using DNAse ...... 60 2.3.3. Reverse transcription and cDNA synthesis ...... 60 2.3.4. Quantitative real-time PCR (qPCR) ...... 61 2.3.5. Visualisation of PCR products by gel electrophoresis...... 62

2.4. Protein Expression Analyses ...... 63 2.4.1. BCA assay...... 63 i) Buffers and Solutions ...... 63 ii) BCA standard curve preparation and loading samples ...... 63 2.4.2. Immunoblotting ...... 63 2.4.3. Human chorionic gonadotropin (hCG) ELISA...... 64 2.4.4. PLAP assay ...... 65

2.5. Immunocytochemical detection ...... 65

2.6. Measurement of caspase activity ...... 66 2.6.1. Caspase 8 ...... 66 2.6.2. Caspase 3/7 ...... 66

2.7. Cell viability (MTT) assay ...... 67

2.8. Transient transfection of genes using siRNA...... 67

2.9. Lipid analysis by mass spectrometry ...... 69 2.9.1. Ceramide and cholesterol Extraction ...... 69 2.9.2. Sphingosine and S1P extraction ...... 71

2.9.3. Standard curves and LC-MS/MS parameters for compounds analyzed using LC-MS/MS...... 72

2.10. Statistics ...... 76

CHAPTER 3. TROPHOBLAST DIFFERENTIATION IN VITRO MODELS ...... 77

3.1. Introduction ...... 77

3.2. Results ...... 78 3.2.1. Biochemical differentiation markers in primary trophoblasts and BeWo cells ..... 78 3.2.2. Caspase activity during syncytialization in term villous trophoblasts ...... 82 3.2.3. Modulating the PKA pathway during trophoblast differentiation ...... 84

3.3. Discussion ...... 86

CHAPTER 4. BCRP TRANSPORTER, SPLS AND TROPHOBLAST DIFFERENTIATION ...... 90

4.1. Introduction ...... 90

4.2. Results ...... 92 4.2.1. Effect of expression knockdown and functional inhibition of BCRP on biochemical trophoblast differentiation ...... 92 4.2.2. Regulation of SPL enzymes and metabolites in response to BCRP silencing ...... 96

4.3. Discussion ...... 99

CHAPTER 5. ROLE OF CERAMIDE BIOSYNTHESIS AND METABOLISM IN THE REGULATION OF TROPHOBLAST DIFFERENTIATION AND SYNCYTIALIZATION ...... 101

5.1. Introduction ...... 101

5.2 Results ...... 102 5.2.1. Expression levels of endogenous aSMase, ceramidase and CERK during spontaneous biochemical trophoblast differentiation and syncytialization...... 102 5.2.2. Regulation of trophoblast differentiation by ceramide and its synthesis/metabolic enzymes ...... 106 5.2.3. Effects of silencing of aSMase and ceramidase expression ...... 115 5.2.4. Downstream signaling targets of ceramide regulating its pro-differentiation actions ...... 119

5.3. Discussion ...... 123

CHAPTER 6. REGULATION OF TROPHOBLAST DIFFERENTIAITON AND SYNCYTIALIZATION BY SPHINGOSINE AND S1P ...... 129

6.1. Introduction ...... 129

6.2. Results ...... 130

6.2.1. Changes in intracellular levels of Sph, secreted S1P and expression of their biosynthesis/metabolic enzymes during spontaneous trophoblast differentiation...... 130 6.2.2. Regulation of trophoblast differentiation by the Sph-S1P pathway...... 134 6.2.3. Effects of silencing SPHK1 on trophoblast differentiation ...... 142 6.2.4. Downstream signaling targets of S1P regulating its anti-differentiation actions...... 144

6.3. Discussion ...... 145

CHAPTER 7. CONCLUSIONS AND FUTURE PERSPECTIVES ...... 150

CHAPTER 8. REFERENCES ...... 157

xii

LIST OF FIGURES

Figure 1.1 Schematic representation of factors involved in trophoblast fusion

Figure 1.2 Structure of sphingolipids

Figure 1.3 Sphingolipid metabolic pathways

Figure 1.4 Transbilayer movement of ceramide

Figure 1.5 Extracellular and intracellular signaling pathways and functions

mediated by S1P

Figure 1.6 Signaling pathways and subsequent cellular responses activated by

ceramide, Sph and S1P

Figure 1.7 Overviewof ABC transporters involved in lipid efflux

Figure 1.8 Effects of TNF-α/IFN-γ on C16, 18, 20 and 24 ceramide accumulation

Figure 2.1 Standard curves for cholesterol, ceramide, Sph and S1P generated

using the Xcalibur software system.

Figure 3.1 Primary trophoblasts isolated from the human term placenta and

visualised using Sigma fastTM 3,3–diaminobenzidine (DAB)

detecting epithelio-specific marker cytokeratin-7

Figure 3.2 Differentiation of primary trophoblasts and BeWo cells in vitro

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Figure 3.3 Caspase 8 and 3/7 activity during trophoblast differentiation in vitro

Figure 3.4 Effects of H89 and FSK on term trophoblast differentiation

Figure 4.1 Suppression of BCRP expression in term trophoblasts using

transient transfection with StealthTM BCRP siRNA

Figure 4.2 Effect of BCRP silencing and inhibition on trophoblasts during

spontaneous differentiation

Figure 4.3 Assessment of BCRP function and trophoblast viability

Figure 4.4 Expression of key SPL hydrolytic enzymes aSMase (A),

ceramidase (B), CERK (C), SPHK1 (D) and S1PP2 (E) in

response to BCRP silencing

Figure 4.5 Effect of BCRP StealthTM siRNA on endogenous levels of

C16 ceramide (A) and Sph (B) measured by LC-MS/MS

Figure 4.6 BCRP expression in response to short chain ceramide (A),

Sph (B) and S1P (C)

Figure 5.1 Expression levels of ceramide synthetic and metabolic enzymes

during trophoblast differentiation in vitro

Figure 5.2 Endogenous levels of ceramide during trophoblast differentiation in vitro

xiv

Figure 5.3 Illustration of pathways targeted with exogenous ceramide

modulators and enzyme inhibitors to investigate their functions

in regulating biochemical and structural trophoblast differentiation

Figure 5.4 Assessment of toxicity of various SPL mediators and inhibitors

in term trophoblasts

Figure 5.5 Regulation of biochemical trophoblast differentiation by

ceramide modulators

Figure 5.6 Differential effect of ceramide modulators on hCG expression vs. secretion

Figure 5.7 Lack of post-differentiation effects of ceramide modulators

Figure 5.8 Ceramide modulators have disparate effects on biochemical

differentiation markers

Figure 5.9 Morphological differentiation in response to ceramide modulators

Figure 5.10 Efficiency of siRNA-mediated transfection in primary trophoblasts

compared to immortalized BeWo cells

Figure 5.11 Silencing of aSMase and ceramidase expression in BeWo cells

using StealthTM siRNA

Figure 5.12 Protein expression of aSMase and ceramidase in FSK-induced

differentiation BeWo cells through 7 days of culture

Figure 5.13 Effect of SP600125, okadaic acid and tautomycin on hCG secretion,

PLAP activity and cell viability by human primary culture of

trophoblasts

Figure 5.14 Phosphorylation of JNK

Figure 5.15 Phosphorylation of Akt

Figure 6.1 Expression of SPHK1 and S1PP2 during trophoblast differentiation

in vitro

Figure 6.2 Levels of intracellular Sph and secreted S1P during trophoblast

differentiation in vitro

Figure 6.3 Illustration of SPLs and enzymes in the Sph-S1P pathway targeted

with modulators and enzyme inhibitors to investigate their role

in trophoblast differentiation and syncytialization

Figure 6.4 Effects of Sph, S1P and SPHK1 inhibitor on trophoblast cell viability

Figure 6.5 Regulation of hCG levels by SPL compounds

Figure 6.6 Effect of DMS on trophoblasts during differentiation

Figure 6.7 Regulation of biochemical differemtiation markers in response

xvi

to SPL modulators

Figure 6.8 Changes in morphological differentiation

Figure 6.9 Efficiency of siRNA-mediated transfection in primary

trophoblasts compared to immortalized BeWo cells

Figure 6.10 Phosphorylation of Akt in response to S1P treatment

Figure 7.1 Role of SPLs in trophoblast differentiation and syncytialization.

xvii

LIST OF TABLES

Table 1.1 Factors that trigger (+) or impair (-) syncytialization of trophoblasts

Table 2.1 List of Reagents

Table 2.2 Final concentrations used of various test substances

Table 2.3 Gene specific primers for quantitative PCR

Table 2.4 StealthTM siRNA duplexes used for transient gene silencing

Table 2.5 Volume added per well in a 24 well plate

Table 2.6 Liquid chromotography conditions

Table 2.7 LC-MS/MS tune parameters for ceramide and cholesterol (A) and

Sph and S1P (B)

Table 2.8 LC-MS/MS instrument methods for C16 Cer, Sph, S1P and their

respective internal controls

xviii

ABBREVIATIONS

Abbreviations Name ABC ATP binding cassette ANOVA Analysis of variance apoA-1 ApolipoproteinA-1 aSMase Acid sphingomyelinase BCA Bicinchoninic acid BCRP Breast cancer resistant protein BrdU 5-bromo2’-deoxy-uridine BSA Bovine serum albumin CAM Cell adhesion molecule cAMP Cyclic adenosine monophosphate CAPS Calcium-dependent activator protein for secretion cDNA Complementary deoxyribonucleic acid CDase Ceramidase Cer Ceramide CERK Ceramide kinase C1P Ceramide-1-phosphate CO2 Carbon dioxide DAB Diamino-benzadine DNA Deoxyribonucleic acid

dNTP Deoxyribonucleotide triphosphate EDG Endothelial differentiation genes EGF Epidermal growth factor EGFR Epidermal growth factor receptor ER Endoplasmic reticulum ERK Extracellular signal regulated kinase FAK Focal adhesion kinase FBS Fetal bovine serum Flt Fms-like tyrosine GCM1 Glial cell missing factor 1 GluCer Glucosylceramide

xix

GM-CSF Granulocyte-macrophage colony stimulating factor GnHR Gonadotropin releasing hormone gp Glycoprotein HAI-1 Hepatocyte growth factor activator inhibitor type-1 hCG Human chorionic gonadotropin HCl Hydrochloric acid HDL High density lipoproteins HEPES 4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid HERV Human endogenous retroviruses HIF Hypoxia-inducible factor H2O2 Hydrogen peroxide hPL Human placental lactogen HRPO Horseradish peroxidase IUGR Intrauterine growth restriction JAK Janus kinase JNK c-Jun N-terminal kinase KCl Potassium chloride LRM Lubrol WX-resistant microdomain LPP Lipid phosphate phosphatases MAPK Mitogen-activated protein kinase Mash Mammalian achaete/scute homologue MDR Multidrug resistant MEK MAPK-ERK kinase MRP Multidrug resistance associated protein mRNA Messenger ribonucleic acid MTT 3-[4,5-dimethyl thiazol-2-yl]-2,5 diphenyl tetrazolium bromide nSMase Neutral sphingomyelinase PA Phosphatidic acid PAP Phosphatide phosphohydrolase PBS Phosphate-buffered saline PC Phosphatidylcholine PE Phosphatidylethanolamine

PFA Paraformaldehyde PG Prostaglandin PIP2 Phosphatidylinositol-4,5-biphosphate PI3K Phosphatidylinositol 3-kinase PKA Protein kinase A PKC Protein kinase C PL Phospholipase PLAP Placental alkaline phosphatase PMA Phorbol 12-myristate 13-acetate PP1 Protein phosphatase 1 PP2A Protein phosphatase 2A PS Phosphatidylserine RNA Ribonucleic acid ROS Reactive oxygen species RT-PCR Reverse transcription-polymerase chain reaction SAPK Stress-activated protein kinase SDK Sphingosine-dependent protein kinase SDS Sodium dodecyl sulphate SEM Standard error of the mean siRNA small interfering RNA SPHK Sphingosine kinase Sph Sphingosine SPHK1 Sphingosine kinase 1 SPL Sphingolipid SM Sphingomyelin S1P Sphingosine-1-phosphate S1PP S1P phosphatase STB Syncytiotrophoblast TAE Tris acetate TGF Transforming growth factor TNF Tumor necrosis factor TNF-R Tumor necrosis factor receptor TRM Triton X100-resistant microdomain

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VEGF Vascular endothelial growth factor

xxii

Introduction

CHAPTER 1. INTRODUCTION

1.1. Trophoblast differentiation and syncytial formation

The placenta is a remarkable organ performing indispensable endocrine, transport and exchange functions throughout pregnancy. It comprises the sole link between the mother and fetus, acting as a selective semi-permeable barrier that facilitates transfer of important nutrients and metabolites while preserving the separation between maternal and fetal blood supplies. It is derived from cells of the embryonic trophectoderm and is essential for the initiation, maintenance and successful conclusion of pregnancy (Cunningham et al. 1989).

The primary cell type of the placenta is the trophoblast, which exhibits a number of different phenotypes and performs a variety of functions according to its location and stage of development. The villous stage of placental development occurs after implantation of the blastocyst, and involves trophoblasts differentiating along two defined pathways: the invasive pathway (extravillous trophoblasts, EVT) and the syncytial pathway involving cytotrophoblasts (CTB, also termed Langerhans cells) and syncytiotrophoblast (collectively termed villous trophoblasts). Extravillous trophoblasts invade the decidua basalis and remodel maternal spiral arteries until they reach the myometrium allowing the placenta to be lodged into the uterus. This is essential to allow an exponential increase in maternal blood flow to serve the demands of the rapidly growing fetus (Baczyk et al. 2009). On the other hand, villous trophoblasts remain attached to the basement membrane, where mononuclear CTBs differentiate and their plasma membranes fuse to form a multinucleated syncytiotrophoblast, also known as the syncytium (Gauster et al. 2009; Malassine et al. 2010).

Trophoblast cell fusion is a finely regulated, dynamic and multifactorial process requiring many biochemical and morphological changes that are currently poorly understood. Formation of the syncytiotrophoblast commences within a week of gestation, and a similar time line is observed in vitro for formation of the syncytial layer in culture. In order to fuse, mitotically active mononuclear CTBs cease proliferating and express pro-fusion genes and proteins. These biochemical and

Introduction molecular changes then allow the cells to recognize and interact with their fusion partners leading to communication initiation between cells and signals exchange resulting in fusion (Malassine et al. 2010). The structural and functional differences that take place during placental development are due to changes in the requirements of the placenta as it matures (Huppertz et al. 1998; Evseenko et al. 2006; Huppertz et al. 2006). The morphological changes observed during trophoblast differentiation are accompanied by alterations in the expression profile of transporters and drug metabolising enzymes (Garland 1998). It is possible that these findings are the underlying cause of the phenotypic differences observed between CTBs and syncytiotrophoblast. As gestation progresses, the relative balance between the CTB and syncytiotrophoblast layers alters as CTBs are lost and the syncytium becomes the predominant layer (Mori et al. 2007). As for the underlying CTBs, several conflicting theories prevail: the more recent theory proposed that during early pregnancy, the continous layer of cuboidal CTBs becomes an incomplete layer comprised of flattened cells with multiple processes used to intercommunicate during later gestation (Jones et al. 2008). An earlier study reported the CTB layer becomes discontinuous over gestation as it cannot keep up with the branching and expansion of the villous surface, albeit a steady increase in cell numbers (Simpson et al. 1992). Another recent study others suggested that the CTB layer becomes thinner but largely maintains its continuity over gestation and is involved in barrier maintenance during late gestation (Mori et al. 2007). It has also been suggested that the rapid growth of the villous area as gestation progresses may be due to the expansion of non-trophoblastic compartments (Mayhew and Simpson 1994). Trophoblast cell cultures provide an in vitro model where isolated mononuclear CTBs can be allowed to aggregate and fuse to form functionally-active, non-proliferative, multinucleated syncytium analogous to that which forms in vivo (Kliman et al. 1986).

While CTBs are considered as trophoblastic stem cells, the syncytium is considered the terminally differentiated layer that forms from fusion of CTBs (Pierce and Midgley 1963). The syncytium comprises of a fetal-facing (basal) and a maternal- facing (apical brush broader) membrane (Ceckova-Novotna et al. 2006). Since the syncytiotrophoblast nuclei do not replicate, it relies solely on fusion of underlying, continuously proliferating CTBs for growth and replenishment throughout (Huppertz and Kingdom 2004; Rote et al. 2010). Together these two cell layers separate fetal

Introduction blood vessels and the mesenchymal connective tissue core from the intervillous space (Mori et al. 2007). Initially it was believed that in order to maintain syncytial integrity and composition, there was continuous disposal of aged nuclei and cytosolic content, in the form of syncytial knots, into the maternal circulation (Huppertz et al. 1998). However, a more recent review concluded there is little evidence to support the concept that turnover of syncytial nuclei takes place in the normal placenta, or that this occurs through an apoptotic-related process (Burton and Jones 2009). They suggested that a proportion of syncytial nuclei are transcriptionally active, that epigenetic modifications underlie the changes in chromatin appearance, and that syncytial nuclei continue to accumulate until term. Moreover, they considered the deportation of trophoblast that has been linked to preeclampsia to be most likely of necrotic origin following ischemic injury (Burton and Jones 2009).

The syncytium, a polarised rate-limiting barrier for transplacental transport, is involved in nutrient and gas exchange and protects the fetus from potential environmental and endogenous toxins; it also synthesizes hormones and steroids essential for fetal growth and development. Syncytial formation is regulated by numerous factors in an autocrine and paracrine manner which can be broadly categorized in to six groups: growth factors, hormones and cytokines, protein kinase and transcription factors, membrane proteins, proteases, physicochemical factors and membrane architecture (Table 1.1) (Morrish et al. 1987; Lyden et al. 1993; Adler et al. 1995; Alsat et al. 1996; Mi et al. 2000; Yu et al. 2002; Frendo et al. 2003; Frendo et al. 2003; Kudo et al. 2003; Yang et al. 2003; Daoud et al. 2005; Knerr et al. 2005; Gauster et al. 2009). However, the syncytialization process is not completely understood and there remains considerable debate regarding the key aspects of biochemical maturation and morphological differentiation of mononuclear cytotrophoblasts into the multinucleated syncytium (Orendi et al. 2010; Rote et al. 2010). Several studies have identified a dissociation between biochemical differentiation/maturation (hCG secretion) and morphological differentiation (fusion/syncytialization) of trophoblasts (Kao et al. 1988; Orendi et al. 2010). In the absence of the latter, the former is still witnessed in serum-free conditions in vitro, indicating that syncytialization is not required for biochemical differentiation although it is an aspect of the differentiation program essential for full barrier and transport function (Kao et al. 1988).

Introduction

Growth factors, EGF + TT Hormones and CSF + TT cytokines

GM-CSF TT +

LIF TT +

TGF-α TT +

VEGF FT, TT +

hCG TT + + PL74 (MIC-1) TT

- TGF-β TT

- TNF-α TT

Protein Kinases ERK1/2 + TT

p38 TT +

PKA B +

Membrane Syncytin 1 + TT, B proteins ASCT1 ? -

ASCT2 + B

CD98 + B

Galectin 3 + B

Connexin 43 + MT

Proteases Caspase 8 + VE

Caspase 10 ? -

Caspase 14 ? -

Introduction

Calpain + TT, B

ADAM12 ? -

Physicochemical Hypoxia - TT, B factors Calcium + TT, B

Membrane PS flip + B architecture

Transcription GCM1 + B factors Mash-2 - B

HIF - FT

LXR - TT

PPARγ +/- TT

Table 1.1. Factors that trigger (+) or impair (-) syncytialization of cultured trophoblasts. Data were obtained from first trimester trophoblasts (FT), midtrimester trophoblasts (MT), term trophoblasts (TT), BeWo cells (B) or villous explants (VE). Some factors (indicated by “?”) have been suggested to play a role in trophoblast fusion, but so far evidence has not been provided (Gauster et al. 2009).

Perturbations in trophoblast differentiation have been associated with faulty syncytial formation causing inefficient nutrient transfer across the placenta, which is associated with numerous pathological clinical conditions including preeclampsia, Down’s syndrome, intrauterine growth restriction (IUGR) and trisomy 21 (Frendo et al. 2000; Yang et al. 2003; Malassine et al. 2010) (Brosens et al. 1977; Himmelmann et al. 1996). Therefore, it is important to better understand the underlying principles of this complex process. While the exact aetiology of preeclampsia remains unclear, this disorder is a leading cause of maternal and perinatal morbidity and mortality (Brown et al. 2005). Recent findings show that IUGR and preeclampsia have distinct pathologies with different trophoblast differentiation profiles (Newhouse et al. 2007). While trophoblast cultures from IUGR placentas show increased trophoblast

Introduction differentiation and syncytial fusion compared to normal cells, trophoblasts from preeclamptic pregnancies exhibit only a moderate increase in syncytialization and no difference in functional differentiation (Newhouse et al. 2007). Contrarily, others have documented evidence of a proliferative CTB phenotype with a reduced syncytial layer in preeclampsia (Alsat et al. 1996). As a consequence of a disruption in the trophoblastic proliferative/differentiating unit during preeclampsia, an increase in apoptosis and turnover of villous trophoblast is observed (Huppertz and Kingdom 2004). Interestingly, pregnancies with both IUGR and preeclampsia exhibit significantly different hormone accumulation and syncytialization profiles compared to those with IUGR only, underscoring the fact that different pathologies can be manifested with differences in trophoblast differentiation and placental development characteristics. In the case of Trisomy 21, trophoblasts aggregate but fuse poorly in vitro, correlating with a downregulation of syncytial hormones, release of hyperglycosylated hCG with diminished bioactivity with a decrease in the number of hCG/luteinising hormone (LH) receptors (Frendo et al. 2000; Massin et al. 2001; Frendo et al. 2004; Pidoux et al. 2004; Pidoux et al. 2007). These studies highlight the importance of hCG and its receptor in trophoblast differentiation and syncytial formation.

1.1.1. Hormones, growth factors and cytokines

The ability of trophoblasts to differentiate and fuse into a multinucleate syncytium and acquire an active endocrine phenotype is under the control of a plethora of hormonal signals (Delidaki et al. 2011). Accurate spatial and temporal expression of placenta- derived hormones that regulate maternal endocrine system and placental functions is crucial for a successful pregnancy (Talamantes and Olgren 1988). The multinucleated syncytium is the source of several hormones that can be measured in abundance in maternal blood in pregnancy (Hoshina et al. 1982). One of the most extensively studied hormones for its role in placental development, in particular villous human trophoblast differentiation, is hCG (Pierce and Parsons 1981). It was initially believed to be synthesized exclusively in the syncytium, but later studies detected traces of hCG mRNA localized to some CTBs, although primarily to the syncytiotrophoblast (Hoshina et al. 1982). Placenta-derived hCG, a member of the gonadotrophin family of glycoprotein hormones, is composed of two noncovalently associated α and β

Introduction subunits. The common α-hCG subunit is encoded by a single a gene (Zimmermann et al. 2003). Unlike α-hCG, β-hCG is encoded by a cluster of six homologous genes localized in chromosome 19 (Fiddes and Goodman 1979; Jameson and Hollenberg 1993). The free α-hCG and β-hCG proteins combine to form an intact biologically active hCG molecule. Synthesis of hCG is in a differentiation-dependent manner, and in turn regulates syncytial formation and self-synthesis via its autocrine and paracrine actions (Shi et al. 1993; Frendo et al. 2003). Secretion of hCG peaks at 9-14 weeks gestation during early pregnancy followed by a rapid decline thereafter, with numerous growth factors contributing towards its synthesis and secretion (Braunstein et al. 1980; Petraglia et al. 1989; Petraglia et al. 1995). The morphological and functional roles of hCG in trophoblast differentiation are temporally regulated by its endogenous and secreted levels, posttranslational modifications and proteolysis in a dose-dependent manner (Birken et al. 1991; Cole et al. 1991; Kardana et al. 1991).

Studies have described various effects of hCG secretion in CTBs, some of which include cell aggregation, expression of a cell adhesion receptor (cadherin) that regulates cell aggregation, expression of the hCG/LH receptor (a G-protein-coupled receptor family member) and synthesis of steroid hormones (Shi et al. 1993). Although the factors regulating hCG release in vivo remain obscure, in vitro studies have shown that calcium (Ca2+) influx into the syncytium via L-type voltage-sensitive Ca2+ channels causes an immediate and sustained release of hCG through a sodium (Na+)-Ca2+ exchange process (Polliotti et al. 1994; Petit and Belisle 1995; Lambot et al. 2005). Expression of the functional cell surface hCG/LH receptor is detected in CTBs and the syncytium (Reshef et al. 1990; Segaloff and Ascoli 1993). Binding of hCG to its receptor activates adenylate cyclase, phospholipase C and ion channels, which in turn control cellular cAMP, inositol phosphates, Ca2+ and other second messengers (Gudermann et al. 1992; Hipkin et al. 1992). Interestingly, while studies have shown that hCG mediates its differentiation effects via the cAMP-protein kinase A (PKA) dependent pathway, inhibiting hCG/LH receptor was found to be ineffective on cAMP-PKA related differentiation effects (Sawai and Azuma 1996; Frendo et al. 2003). Furthermore, activation of cAMP-PKA inducing trophoblast differentiation and fusion has been shown independent to hCG secretion, most likely by targeting other differentiation specific genes (Yang et al. 2003; Lambot et al. 2005; Orendi et al. 2010). In line with these findings, Orendi et al. have recently presented data

Introduction suggesting regulation of syncytialization independent of hCG secretion. A new model for villous trophoblast syncytial formation has also recently been proposed indicating differentiation signals may trigger pathways leading to hCG secretion resulting in biochemical aspects of differentiation, without directly affecting intercellular fusion (Rote et al. 2010).

Production of hCG regulates and is regulated by steroid hormones that play a particular relevance in placental function, for example progesterone, estrogens and placental gonadotropin-releasing hormone (GnRH) (Khodr and Siler-Khodr 1978; Siler-Khodr and Khodr 1981; Petraglia et al. 1987). Release of progesterone from placental cells in response to hCG in turn inhibits GnRH-stimulated release of hCG from cultured trophoblasts, whereas estrogen has a positive effect on GnRH- stimulated hCG secretion and mRNA levels (Wilson et al. 1984; Petraglia et al. 1989; Ringler et al. 1989). The increased synthesis of progesterone is also positively regulated by estrogen and correlates with syncytial formation (Chaudhary et al. 1992; Petraglia et al. 1995). In addition, other trophoblast glycoprotein hormones such as inhibin and activin have an inhibitory and stimulatory effect on the GnRH-hCG axis, respectively (Keelan et al. 1994). Colocalization of these peptides and hormones in trophoblasts support the hypothesis of autocrine/paracrine steroid-protein interactions in placental development (Petraglia et al. 1995).

Another major placental hormone that is associated with syncytialization is human placental lactogen (hPL) (Alsat et al. 1996). Hormonal signals finely tune specific mechanisms that stimulate expression of a specific cassette of important fusogenic genes such as syncytin-1 and -2 (discussed below) and increase production of hormones such as hCG and hPL (Knerr et al. 2005). Unlike hCG, hPL mRNA is found only in the syncytial layer with its secretion levels steadily increasing and reaching a plateau at 36 weeks of gestation, and then declining slightly after the 37th week (Letchworth et al. 1978; Geyer and Nothnagel 1984). While hPL serum levels depend on the mass of trophoblast and on differentiation and synthesis in the syncytiotrophoblast, the peak of serum hCG levels as well as the maximum immunohistochemical staining for this hormone occurs at 9-14 weeks gestation as mentioned above, a time when the mass of trophoblast is relatively small. The attainment of the highest maternal serum hCG levels at a time when the trophoblast mass is small implies maximum production and synthesis of hCG in the first trimester

Introduction

(Beck et al. 1986). In vitro, levels of hPL and hCG peak at 24 hr and 72 hr of cell culture during trophoblast fusion, respectively (Wich et al. 2009).

There are a number of growth factors and cytokines that have been extensively studied for their stimulatory roles in regulating syncytialization. Some of the more widely studied ones include epidermal growth factor (EGF), granulocyte-macrophage colony stimulating factor (GM-CSF), transforming growth factor (TGF)-α and vascular endothelial growth factor (VEGF) (Morrish et al. 1987; Garcia-Lloret et al. 1994; Crocker et al. 2001; Yang et al. 2003). On the other hand, inhibition of syncytial formation has been equally documented in response to tumor necrosis factor (TNF)-α and TGF-β (Morrish et al. 1991; Leisser et al. 2006). Interestingly, while some studies show leukemia inhibitory factor (LIF) upregulating syncytial formation, others have shown it having a negative impact on the process (Nachtigall et al. 1996; Yang et al. 2003). All together, aberrant concentrations of growth factors, cytokines and receptor isoforms affect trophoblast turnover by influencing downstream signaling (Gauster et al. 2009).

1.1.2. Membrane proteins

CTB aggregation is a Ca2+- and protein synthesis-dependent process; therefore, it is logical to postulate that Ca2+-dependent membrane proteins play an essential role in cell adhesion required for normal trophoblast differentiation (Coutifaris et al. 1991). These cell aggregates establish extensive interactions with one another through the formation of tight junctions, gap junctions, ephrins, cadherins, catenins, and ADAMs (a disintegrin and metalloproteases) indicating a multiplicity of adhesive interactions (Douglas and King 1990; Aplin et al. 2009).

One of the major membrane protein families associated with trophoblast fusion is the connexin family, in particular connexin 43. Gap junctions contain transmembrane channels composed of connexin dodecamers (Aplin et al. 2009). These transmembrane channels allow passage of small signaling molecules, such as cAMP and Ca2+, that regulate differentiation and fusion (Malassine et al. 2010). Connexin 43 is found at the CTB-CTB interface, as well as the CTB-syncytium interface, and its expression increases during trophoblast syncytialization (Frendo et al. 2003;

Introduction

Malassine and Cronier 2005). It has been suggested that gap junction communications may be a prerequisite of villous trophoblast differentiation since connexin-deficient animal models presented altered trophoblast structurally and functionally (Brown et al. 2005). Studies have found a number of secondary membrane proteins linked to these gap junctions, including phosphatases, kinases, scaffolding proteins, chaperones and cell signaling molecules (Herve et al. 2004). Amongst these, zona occludens-1 (ZO-1; a tight junction protein) shows colocalization with connexin 43 at intercellular boundaries of aggregated CTBs and the CTB-syncytium interface during differentiation, with disrupted membrane fusion following silenced ZO-1 expression (Malassine et al. 2010). In addition, ZO-1 expression has been detected in the syncytial microvillous membrane. Interestingly, as connexin 43 expression is upregulated during syncytialization, the expression of ZO-1 decreases during this process (Aplin et al. 2009).

Cadherins represent another significant family of membrane proteins associated with trophoblast differentiation and syncytial fusion, and can be broadly divided into four subgroups: the classical (type I) and closely related type II cadherins, desmosomal cadherins and protocadherins (Aplin et al. 2009). Of the classical and non-classical cadherins, the most commonly studied cadherins in trophoblast differentiation include E-cadherin and cadherin-11, respectively (Aplin et al. 2009). Cadherins belong to a superfamily of integral glycoproteins that facilitate cell-cell adhesion between epithelial cells in a calcium-dependent manner (Gallin 1998). They are able to mediate morphogenesis and cell-cell interactions by interacting with at least three cytoskeletal- associated proteins: α-, β- and γ-catenins (Edelman 1989; Ozawa et al. 1989). Catenins, in turn, link cadherins to the underlying cytoskeleton and are involved in activating several downstream signaling pathways. β-catenin, in particular, plays a significant role in the Wnt pathway, which is required for trophoblast differentiation (Pollheimer et al. 2006).

Cadherins are differentially expressed during the aggregation, differentiation and fusion of trophoblasts, governing cell fate and subsequent tissue formation. This is mediated partly by intracellular signaling via the cadherin-catenin complex (Gumbiner 1995; Dale 1998; Getsios et al. 2000). E-cadherin expression in isolated trophoblast cultures shows levels peaking 24 h post isolation, temporally coinciding with maximal trophoblastic cell aggregation, after which the cells begin to undergo differentiation

Introduction and fusion, accompanied by a decline in membrane-bound E-cadherin in vitro and in vivo (Kliman et al. 1986; Getsios et al. 2000). This trend is mimicked by β-catenin expression, which has a mixed membranous and granular cytoplasmic distribution, in differentiating villous trophoblasts (Getsios et al. 2000).

Although the precise role of the E-cadherin/β-catenin complex in trophoblast turnover is still poorly defined, the loss of E-cadherin and β-catenin expression suggests that their functional significance may contribute towards morphological trophoblast differentiation by altering many characteristics of these cells, including integrity and organization of intracellular junctional complexes (Douglas and King 1990; Coutifaris et al. 1991). In addition, downregulation of the E-cadherin/β-catenin complex expression may also regulate remodeling of the cytoskeleton and expression of transcription factors, either individually or in combination, thereby influencing cell differentiation (Douglas and King 1993; Dakour et al. 1999).

While E-cadherin expression is generally lost as cells fuse, some studies have witnessed weak staining within the syncytiotrophoblast cytoplasm and limited redistribution to the basal membrane (MacCalman et al. 1996). A plausible explanation for this may be that either CTB-derived E-cadherin requires time to breakdown on entry to the syncytium, or that its redistribution to a limited extent maintains heterotypic adhesion between the syncytiotrophoblast and CTBs (Brown et al. 2005). Alterations in the expression of E-cadherin/β-catenin complex may play a role in normal and pathological placental development, with a downregualtion of the complex observed in gestational trophoblastic diseases and the opposite detected in preeclampsia (Li et al. 2003).

On the other hand, cadherin-11 expression increases during formation of the multinucleated syncytium, accompanied by augmented mRNA expression levels of the β-hCG (Getsios et al. 2000). Another commonly studied membrane protein desmoplakin, similar to E-cadherin, has been used as a morphological marker of villous trophoblast differentiation since it is downregulated as trophoblasts undergo differentiation and syncytialization (Douglas and King 1990; Aplin et al. 2009).

Human endogenous retroviruses (HERV) have been increasingly studied in the last decade in trophoblast research. They comprise 8% of the human genome, with 16

Introduction functional non-defective genes identified, of which two are highly and specifically expressed in the placenta. These genes encode HERV-W Env glycoprotein (syncytin- 1) and HERV-FRD Env glycoprotein (syncytin 2) (Blond et al. 2000; Lander et al. 2001; Blaise et al. 2003). During trophoblast syncytial formation, cAMP-induced phosphorylation of PKA leads to stimulation of transcription factor glial cells missing 1 (GCM1), a placenta-specific transcription factor, which induces syncytin-1 generation and eventually trophoblasts fusion, accompanied by elevated hCG production (Knerr et al. 2005; Orendi et al.). Induction of trophoblast differentiation in vitro is associated with an upregulation in syncytin-1 expression, which is exclusively found in the syncytiotrophoblast (Frendo et al. 2003). However, studies have reported a progressive decrease in syncytin-1 and GCM1 transcripts following the initial upregulation after 48 hr of cell culture, perhaps indicating a regulatory limitation of the fusion process (Wich et al. 2009).

In addition, detection of syncytin-1 in non-fusogenic extravillous trophoblast suggested a non-central role in syncytial fusion. Furthermore, downregulation of syncytin-1 receptor an amino acid transporter Bo (ASCT2) activity, expressed at the syncytial basal membrane, is also observed as cells syncytialize. Nevertheless, expression of syncytin is spatio-temporally regulated in a highly specific manner to maintain an integral and functional syncytiotrophoblast layer (Yu et al. 2002). The precise function of syncytin-2 in trophoblasts, on the other hand, remains to be elucidated, although its expression declines as trophoblasts undergo differentiation and fusion. (Malassine et al. 2010).

Another membrane protein that has been studied with respect to the formation of the multinuclear syncytium in vitro is placental alkaline phosphatase (PLAP) (Bax et al. 1989; Bullen et al. 1990). This glycosylphosphatidylinositol (GPI)-anchored sialglycoprotein is expressed in large amounts in the syncytium; however, controversies exist regarding its localization in CTB. Some studies have shown low levels of expression in CTBs, thereby questioning its use as a morphological differentiation marker (Leitner et al. 2001). Others show it is exclusively expressed in the syncytiotrophoblast and argue that any traces of this enzyme in CTB cultures represents remnant syncytial fragments produced during isolation (Guilbert and Winkler-Lowen 2007). These discrepancies are most likely due to differences in isolation techniques and culture conditions, which in turn have a marked effect on cell

Introduction purity and trophoblast differentiation (Bloxam et al. 1997). Nevertheless, its expression and activity are still highly upregulated as trophoblasts differentiate and fuse (Leitner et al. 2001).

1.1.3. Protein kinase

PKA-dependent signaling was one of the first downstream pathways to be linked with regulation of trophoblast differentiation (Lohmann and Walter 1984; Keryer et al. 1998). Detailed studies on this kinase revealed a central role in biochemical and morphological trophoblast differentiation. PKA is activated by cAMP, a second messenger that is produced during cell aggregation and syncytialization and is directly associated with regulation of this process (Keryer et al. 1998). In addition to PKA, other well established downstream signaling targets that mediate syncytialization include extracellular signal activated kinase (ERK)1/2 and p38 mitogen-activated protein kinases (Daoud et al. 2005). Both these mitogen-activated protein kinases (MAPK) play central roles in regulation of cell differentiation in various cellular systems, including chondroblasts, erythroblasts, myoblasts and neurones (Nebreda and Porras 2000; Kohmura et al. 2004; Lee et al. 2004). A drastic decrease in expression of ERK1/2 and p38 in isolated trophoblasts with increasing days of culture has been reported, implying a role in initiating trophoblast differentiation (Daoud et al. 2005). Suppressing the activity of these proteins led to impaired differentiation, with a greater effect seen in response to p38 compared to ERK1/2, suggesting a more significant role of this kinase. Furthermore, potential cross-talk between these pathways during differentiation may be likely, since inhibition of one results in cross activation of the other. Interestingly, this compensatory mechanism on its own is incapable of triggering initiation of trophoblast differentiation (Daoud et al. 2005).

1.1.4. Proteases

Caspases (aspartate-specific cysteine proteases) are mainly known for their key role in programmed cell death, but have recently attracted interest with respect to cell differentiation. Examples for caspase-mediated differentiation exist and are quite

Introduction diverse (Gauster et al. 2009). Previous studies have identied a role of caspases in haematopoiesis, including erythrocyte maturation and macrophage differentiation as well as platelet formation (Droin et al. 2008). They are crucial in regulating cell differentiation in various other systems and thus, it is not surprising that caspases have been associated with differentiation of villous CTBs (Weil et al. 1999; Eckhart et al. 2000; Ryan and Salvesen 2003; Zandy et al. 2005; Murray et al. 2008). Although there is extensive literature on the unique differentiation and fusion properties of mononuclear CTBs, including multiple studies on numerous factors regulating this process, the precise role of the cysteine-aspartic proenzyme caspase 8 remains highly controversial.

Formerly, it was believed that during initial differentiation of CTBs to form the syncytiotrophoblast, early and reversible stages of the apoptotic cascade were involved, including cleavage of the caspase 8 substrate alpha-fodrin and the translocation of phosphatidylserine (PS) from the inner to outer leaflet of the plasma membrane (Huppertz et al. 1998; Huppertz et al. 1999). Further, it was widely accepted that these pre-apoptotic stages were prerequisites for syncytial formation, since blocking caspase-8 activity (detectable in CTB cells prior to fusion) prevents PS externalization and subsequent syncytialization (Adler et al. 1995; Vogt et al. 1997; Huppertz et al. 1999; Black et al. 2004; Huppertz and Kingdom 2004; Leslie et al. 2005). However, a recent review highlighted a number of discrepancies in the data supporting the existing dogma, and proposed a new trophoblast differentiation model based on more recent findings (Rote et al. 2010). The authors proposed that the caspase 8 holoenzyme and the proteolytically active form may potentially have different roles, with the former contributing to early differentiation and the latter cleaving fodrin at the last steps of fusion. Suppression of caspase 8 expression and/or activity confirmed potential differentiation-related roles, unrelated to initiation of syncytial fusion. It should be appreciated that the basis of the association between hCG production, cell cycle departure, expression of fusion proteins, PS efflux and cytoskeletal rearrangement during villous trophoblast differentiation and intercellular fusion remains unclear. Further studies are needed to resolve whether any aspect of the model accurately depicts in situ placental development (Rote et al. 2010).

Interestingly, there are also conflicting theories on the role of downstream effector caspases 3 and 6 in differentiation. Studies have shown these caspases are inactive in

Introduction differentiating CTB (Huppertz et al. 1998; Huppertz et al. 1999), with activation of their inactive proforms only occurring once the cells are terminally differentiated (Black et al. 2004). However, Yusuf et al. reported that caspase 3 activity declined between 24 h to 72 h post-isolation, and that activation of these caspases is triggered while CTBs are in their mononuclear state; subsequent intercellular differentiation and fusion progresses in the absence of further caspase activation (Yusuf et al. 2002). Research lacks consensus on the non-apoptotic role of caspases in differentiation and cell division, mainly due to considerable variablility in the specific role of caspases among different cell types (Maelfait and Beyaert 2008).

1.1.5. Physicochemical factors

Placental development and consequently fetal growth are also strictly regulated by physiological factors. Oxygen tension, being a function of uterine blood flow, plays a vital role in CTB proliferation and guiding differentiation to the invasive phenotype (Red-Horse et al. 2004). Early stages of the formation of feto-maternal interactions occur in relatively hypoxic (reduced oxygen tension) conditions, due to plugging of the spiral arteries by endovascular trophoblasts resulting in a limited amount of uterine blood flow to the conceptus (Rodesch et al. 1992). Trophoblasts display a predominantly proliferative phenotype under low oxygen levels (e.g. ≤ 3%), comparable to in vivo conditions during initial stages of gestation (Soleymanlou et al. 2005). This hypoxic environment during the first trimester is essential for successful pregnancy and favours CTB proliferation over differentiation along the invasive pathway. However, subsequent studies related extreme hypoxia to impaired trophoblast differentiation and syncytialization (Alsat et al. 1996; Kudo et al. 2003). Studies in forskolin (FSK)-induced differentiating BeWo cells report diminished expression and function of syncytin and its receptor ASCT2 upon exposure to low ambient oxygen (2-9% oxygen; hypoxia) compared to control cells (20% oxygen; normoxia) (Kudo et al. 2003). Consequently, this suppressed the normal process of cell fusion necessary for syncytial formation and contributed to syncytiotrophoblast abnormalities characteristic of preeclampsia.

Introduction

These findings were replicated in primary trophoblast cultures showing failure of aggregated CTB to fuse into syncytiotrophoblast under hypoxic conditions, and the effects were reversible when cells were reoxygenated (Alsat et al. 1996). During hypoxia, E-cadherin and desmoplakin expression levels in CTBs were not downregulated as expected upon cells fusion; however, diminished hCG production was seen compared to controls (Alsat et al. 1996). These studies highlight the relevance of physiochemical factors in regulation of trophoblast differentiation and syncytialization. The formation of a functional syncytiotrophoblast layer is impaired during hypoxia in vitro due to a defect in the CTB fusion process, providing a plausible explanation for the higher number of CTBs and a reduced syncytial layer observed in placentas from pathological pregnancies (Alsat et al. 1996).

1.1.6. Membrane architecture

The perception of the plasma membrane as a homogenous phospholipid bilayer is long outmoded and has been replaced by concepts of lateral, structural and compositional heterogeneity manifesting as membrane microdomains of variable size, composition and function in a state of constant flux (Ikonen and Parton 2000; Cremesti et al. 2002; Nabi and Le 2003; Parton and Richards 2003; Pike 2006). Extensive studies have attempted to define the properties and functions of these specialized microdomains (historically called lipid rafts) (Pike 2004; Radeva et al. 2005; Pike 2006). Several different definitions of membrane microdomains, their composition and structure exist based on various methods of isolation. However, they were eventually collectively defined as “small (10-200 nm), heterogeneous, highly dynamic, sterol- and sphingolipid-enriched domains that compartmentalize cellular processes, and at times stabilize to form larger platforms through protein-protein and protein-lipid interactions” (Pike 2006). Terms that have widely been used to describe membrane microdomains such as “lipid” rafts, “detergent resistant”, “plasma membrane” and “liquid-ordered” are now no longer considered particularly accurate (Pike 2006), although are still used in light of the lack of alternatives.

The cell membrane comprises of three main classes of lipids: glycerophosopholipids, sterols and (sphingolipids) SPLs. PS, the most abundant phospholipid present in the

Introduction inner layer of the plasma membrane, is generally maintained in a tightly-regulated asymmetrical distribution; it is flipped to the outer leaflet prior to syncytial formation (Black et al. 2004; Huppertz and Kingdom 2004). Given that PS efflux is an early characteristic of apoptosis and dependent on caspase activation, and is considered essential for syncytial formation, trophoblast differentiation was postulated to be dependent on an early apoptotic processes. However, recent findings concluded PS efflux in CTB was unrelated to apoptosis and was reliant on transporters that are adenosine triphoshate (ATP)- and PKA-dependent (Rote et al. 2010). In fact, intercellular fusion seen in other cellular systems, for example myoblasts and between sperm and egg, are also dependent on PS efflux, thereby suggesting it may be a universal and obligatory component of the process (Rote et al. 2010).

SPLs are primarily found in the external leaflet of the plasma membrane (Sietsma et al. 2001). These lipids, distributed asymmetrically across the plasma membrane, are translocated from one leaflet of the bilayer to the other via the “flip-flop” mechanism. The movement of lipids from the extracellular to the inner leaflet of the plasma membrane is termed as “flip” and translocation in the opposite direction is “flop” (Ikeda et al. 2006). In the plasma membrane, SPLs are key players in the stability and formation of membrane microdomains which are enriched in SPLs and cholesterol (Munro 2003). These low density plasma membrane microdomains are a result of hydrophobic interactions via tight hydrogen-bonding between SPLs and cholesterol that exist as lateral assemblies within the plasma membrane (Cremesti et al. 2002). This tight packing prevents SPLs (e.g. ceramide) to transfer spontaneously between lipid bilayers, with decreased flip-flop activity (Blitterswijk et al. 2003). In addition to the plasma membrane, localization of SPLs in the mitochondria, Golgi apparatus, endosomes, lysosomes, ER and nucleus has been observed (Futerman and Hannun 2004).

SPLs were initially considered predominantly for their structural role as mere building blocks of biological membranes (Bartke and Hannun 2009). Over time, these compounds have been given their due importance as key bioactive molecules playing central roles in regulating various signal transduction pathways. Induction of bioactive SPLs by several stimuli results in triggering a range of downstream targets that mediate their various effects of cell function (Bartke and Hannun 2009). Although there is a large body of literature describing the regulation of cell differentiation by

Introduction these lipids in various cellular systems, surprisingly, there is very little known with regards to their role in trophoblast differentiation.

1.1.7. Transcription factors

During trophoblast differentiation, stimulation of the cAMP-PKA pathway leads to activation of downstream GCM1, as mentioned earlier. The first transcription factor discovered to play a role in trophoblast differentiation and fusion, GCM1, triggers synthesis of membrane fusion protein syncytin-1 (Wich et al. 2009). GCM1 has two PKA phosphorylation sites that are activated leading to enhanced expression of syncytin-1 (Pidoux et al. 2007). However, reports suggest it is likely that the regulation of syncytin-1 expression by GCM1 may be cell type-dependent, or that syncytin-1 may also be regulated by other placenta-specific factors, since elevated GCM1 levels in HeLa cells failed to induce any syncytin-1 expression (Yu et al. 2002). GCM1 is a key player in regulating the balance between trophoblast proliferation and differentiation, a crucial step for normal placental development (Baczyk et al. 2009). Diminished GCM1 expression in first trimester villous trophoblasts has been associated with impaired syncytiotrophoblast formation through lack of syncytial fusion, accompanied by an enhanced proliferating CTB layer. Furthermore, formation of the syncytium was completely prevented in denuded villous explants cultures by GCM1 silencing (Baczyk et al. 2009). It is worth noting that severe early preeclampsia presents with downregulated GCM1 levels, in accordance with the reduced placental syncytialization (Chen et al. 2004). These findings underscore the significance of this transcription factor in villous trophoblast differentiation.

The most commonly studied oxygen-sensing transcription factor in the context of regulation of trophoblast fusion is hypoxia-inducible factor-1 (HIF-1). This regulates expression of genes such as VEGF, mammalian achaete/scute homologue (Mash)-2 and soluble fms-like tyrosine kinase (sFlt)-1 (Wang and Semenza 1993; Jiang et al. 2000; Nevo et al. 2006). Upregulation of Mash-2 is observed during hypoxia, and its overexpression results in inhibited syncytial formation (Jiang et al. 2000). Similarly, serum-bound sFlt-1 captures pro-differentiation growth factor VEGF, thereby impairing its regulatory actions and subsequently downregulating syncytialization

Introduction

(Nevo et al. 2006). In preeclamptic women, overexpression of HIF-1 and TNF-β is observed which is indicative of altered trophoblast differentiation and fusion processes (Rajakumar et al. 2003; Nishi et al. 2004).

Other identified transcription factors regulating trophoblast differentiation include nuclear receptors, liver x-receptors (LXR) and peroxisome proliferator activated receptors (PPAR) (Schaiff et al. 2000; Fournier et al. 2007; Aye et al. 2011). LXRs, activated by oxysterols, have an inhibitory effect on trophoblast differentiation (Aye et al. 2011). On the other hand, PPARγ ligands modulate trophoblast differentiation in a ligand-dependent manner. Activation of PPARγ with troglitazone promotes trophoblast differentiation as shown by increased hCG secretion, whereas the endogenous agonist 15d-PGJ2 prevents this process (Schaiff et al. 2000).

1.1.8 Summary

To summarise, fusion of mononuclear CTBs to form the multinucleate syncytium is regulated by multiple factors process (Figure 1.1). It is initiated by the actions of growth factors, cytokines and hormones from the fetal and/or maternal environment binding to their cognate receptors on the plasma membrane of villous CTBs. This results in activation of downstream second messenger cascades such as those involving PKA, ERK1/2 and p38, causing upregulation in protein expression of factors such as GCM1, which in turn drives transcription of fusogenic factors. Phases 1 to 5 in Figure 1.1 illustrate a simplified overview of membrane proteins binding to their respective receptors resulting in remodelling of the cytoskeleton.

Introduction

GCM1

Figure 1.1. Schematic representation of some key factors involved in trophoblast fusion (Gauster et al. 2009).

1.2. Sphingolipids

SPLs were first discovered by Thudichum in 1876, and for nearly 100 years there was very little known about these compounds besides their structural role in cell membranes (Bartke and Hannun 2009). Interest in this family of highly bioactive compounds was rekindled in the 1980s and an increasing list of biological effects was ascribed to them. This list now includes numerous cellular activities, including differentiation, proliferation, death, migration, cell-cell and cell-matrix interactions, intracellular and extracellular signaling and maintenance of lipoprotein and membrane structure; most of these functions are structure-specific (Hakomori 2003). SPLs comprise of long chain alkanes or alkenes, also known as ‘sphingoid’ base backbones (hydrophobic region), which serve as the central moiety of these compounds (Figure 1.2). Free sphingoid bases have a core structure of approximately 14-20 carbons in length, hydroxyl-substituents at positions 1 and 3, and an amino group at position 2, which encompass a wide array of sphingosines (Sph), sphinganines and phytosphingosines (Merrill et al. 2005). The core structure may vary in the length of the alkyl chain (usually 18 carbons long in mammals), the number of double bonds, an additional hydroxyl group at position 4 and branching methyl groups (Merrill et al.

Introduction

2005). Formation of ‘complex SPLs’ [for example ceramide, sphingomyelin (SM) and glucosylceramides (GluCer)] occurs as a consequence of conjugation at the 1- hydroxyl position with a polar group and derivatization on the 2-amino group with a long chain fatty acid (Merrill et al. 2005) (Figure 1.2). The numerous structural possibilities as a result of various combinations of hydrophobic backbones and headgroups is bewildering, rendering these compounds extremely complex and diverse (Merrill and Sandhoff 2002).

Figure 1.2. Structure of sphingolipids. For simplicity, only one kind of sphingoid base (sphingosine, in blue) is shown to which only one kind of fatty acid (palmitic acid, in red) is N-acylated. Similarly, only one glycosphingolipid (GSL), glucosylceramide (black), is shown, but sequential addition of other carbohydrate residues results in >500 known GSL structures (Futerman and Hannun 2004).

SPL biosynthesis and metabolism takes place in multiple subcellular locations (Futerman and Hannun 2004). A major challenge in the research of SPLs is to explain how the multiple SPL-dependent signaling events are modulated, and how they are spatially and temporally regulated according to subcellular localization (Futerman and Hannun 2004). Moreover, the complexity of sphingolipid metabolism is accentuated by the number of enzymes involved in these pathways and their roles in mediating

Introduction distant and often paradoxical effects (Reiss et al. 2004). The SPLs that have been most widely studied regarding their roles in regulating cell differentiation and are also most relevant to this thesis are ceramide, sphingosine (Sph) and sphingosine-1-phosphate (S1P). Therefore, these three SPLs are the focus of this thesis (Kolesnick and Kronke 1998; Hannun and Luberto 2000).

1.2.1. Ceramide i) Synthesis and Metabolism

Ceramide, an N-acylsphingosine, is a ubiquitous SPL at the heart of a complex web of synthetic and metabolic pathways and the hydrophobic backbone of complex SPLs such as SM and GluCer (Cremesti et al. 2002) (Sweeney et al. 1998; Pettus et al. 2002; Rotolo et al. 2005). It has a minimal hydrophilic region comprising two hydroxyl groups, and also serves as a precursor of other SPL compounds (Futerman and Hannun 2004). Different ceramide species arise as a result of fatty acid chains with varying saturation, hyrdroxylation and length (from 2 to 28 carbons; C16 and C28 being the most abundant in mammals) (Cremesti et al. 2002). Different genes from the same family determine the specificity of N-acylation substrate, resulting in the generation of different ceramide species.

Ceramide analogues with differential chain lengths have distinct structural and functional identities in different tissues due to tissue-specific expression of these genes (Venkataraman et al. 2002; Riebeling et al. 2003). Accordingly, the use of exogenous ceramide to investigate its biological effects may not always be an ideal model to simulate physiological responses. This is because the exogenously added short chain molecules may redistribute in a non-physiological manner within cell compartments, thereby leading to misinterpretation of results (Gangoiti et al. 2010). However, due to the highly hydrophobic nature of long chain ceramides, their short chain counterparts are widely used as an alternative for in vitro studies. Distributed primarily in Golgi, endoplasmic reticulum (ER) and plasma membrane, the intracellular location of ceramides influences its fate and activity (Cremesti et al. 2002; Blitterswijk et al. 2003). Ceramide levels are determined primarily by the enzymes that synthesize and catabolize ceramide (Pettus et al. 2002).

Introduction

The key pathways involved in ceramide synthesis include the de novo pathway (mediated by ceramide synthase (CerS)), the sphingomyelinase (SMase)-mediated pathway (via acid or neutral SMase), and catabolism of complex glycosphingolipids with eventual breakdown into Sph, which can be recycled via reacylation to produce ceramide through the salvage pathway (Futerman and Hannun 2004; Kitatani et al. 2008). The enzymes involved in the clearance of ceramide include SM synthase, ceramidase, ceramide kinase (CERK) and glucosylceramide synthase (GCS), which act by converting ceramide to SM, Sph, ceramide-1-phosphate (C1P) and GluCer, respectively (Luberto and Hannun 1998; Liu et al. 1999; Strelow et al. 2000). Generation of Sph from ceramidases, further serves as a precursor for synthesizing S1P via sphingosine kinase (SPHK) activity (Futerman and Hannun 2004). See Figure 1.3.

The de novo pathway commences in the endoplasmic reticulum (ER) with serine and palmitoyl CoA being metabolized through a series of steps to dihydrosphingosine (sphinganine) that undergoes acylation by CerS to generate ceramide; this is then rapidly translocated to the Golgi apparatus where it is metabolized to glycosphingolipids, C1P and SM. CerS has six isoforms with substrate preferences for specific chain length fatty acyl CoAs, thereby generating ceramides with distinct acyl- chain lengths (Kitatani et al. 2008). Due to its hydrophobic nature, ceramide is transported across the cytosol to the Golgi by either vesicular or non-vesicular transport (Fukasawa et al. 1999). Hanada et al. have isolated and characterized a protein called CERT, which extracts ceramide from the plasma membrane and targets it to the Golgi apparatus (Hanada et al. 2003). It may be possible that similar pathways are employed to deliver precursors to the mitochondria or nucleus. Subsequently, the glycosphingolipids and SM generated in the Golgi apparatus (the major site of synthesis) are translocated to the plasma membrane microdomains via vesicular traffic. SM on both sides of the plasma membrane is further hydrolysed to produce ceramide via the SMase pathway (Figure 1.3).

SMases belong to the phosphodiesterase superfamily; they catalyze the conversion of SM to ceramide and phosphocholine, the latter being an inert metabolite (Hannun 1996; Hannun and Obeid 2002). Three distinct forms of SMases have been identified in vitro based on their pH optima: acidic, neutral and alkaline SMases. Interestingly, ceramides produced by different pools of SM on either side of the plasma membrane

Introduction at different times in response to cell stimulation have distinct functions (Blitterswijk et al. 2003). Rapid ceramide synthesis in response to death receptor ligation at the plasma membrane by hydrolysis of SM by acid SMase (aSMase) plays more of a membrane structural role, involving microdomain fusion, capping, membrane curvature and triggering various downstream signals. On the other hand, slow ceramide synthesis by neutral SMase (nSMase) in the membrane is involved in induction of apoptosis, while ceramide produced by the de novo pathway is mainly involved in synthesis of complex SPLs, assembly of lipid rafts and vesicular trafficking. Ceramide can also enter the mitochondria, where it plays a key role in regulating death signals via retrograde vesicular trafficking from SM at the plasma membrane during apoptosis, or via sites of close contact with the ER (Daum and Vance 1997; Prinz et al. 2000; Marsh et al. 2001). In addition, due to its ‘channel forming’ actions, the minor pool of mitochondrial ceramide may exert a membrane structural role (Blitterswijk et al. 2003).

Spatial separation of enzymes allows differential activation of these distinct pathways associated with ceramide generation. However, it remains unclear how signaling pools of ceramide are separated from metabolic pools within a cell. Obviously, compartmentalization is an essential aspect for ceramide function (Hannun and Obeid 2002). Metabolism of SPLs in response to extracellular agents is highly complex and can be specific to a cell type, or regulated by signals produced by allosteric mechanisms, post-translational modifications or enzyme expression patterns (Futerman and Hannun 2004). Accordingly, cellular ceramide homeostasis is governed by which ceramide-generating pathway is activated and its spatial-temporal properties (Kitatani et al. 2008). Being metabolically juxtaposed, the regulation of the metabolism of these lipid mediators is of utmost importance in determining cell fate (Taha et al. 2006). Figure 1.3 gives a brief overview of the complex synthesis and degradation pathways of SPLs.

Introduction

Figure 1.3. Sphingolipid metabolic pathways. Metabolic pathways for ceramide synthesis composed of the sphingomyelinase pathway, the de novo pathway, the exogenous ceramide-recycling pathway, and the salvage pathway. Dotted lines indicate the pathway of ceramide synthesis resulting from recycling/salvaging sphingosine (Kitatani et al. 2008).

Dysregulation of SPL metabolism leads to the establishment and progression of a variety of diseases, including neurodegenerative diseases such as Alzheimer’s or Parkinson’s disease, cardiovascular diseases, chronic inflammation, or cancer (Merrill and Jones 1990; Merrill 2002). In addition, genetic defects in the metabolic enzymes involved in breakdown of SPLs in acidic late endosomes/lysosomes during the salvage pathway leads to lysosomal accumulation of the substrate lipids; this then causes a group of disorders termed “sphingolipidoses”, examples of which include Farber’s disease, Niemann-Pick type diseases A and B, and Gaucher disease (Park and Schuchman 2006; Eliyahu et al. 2007; Kitatani et al. 2008; Bartke and Hannun 2009). SPL metabolizing enzymes are targets for development of novel therapeutic compounds for treatment of these illnesses.

Introduction ii) Structural role

A major role of ceramide has also been implicated in membrane fusion, pore formation and generation of lipid microdomains in the plasma membrane (Cremesti et al. 2002). As mentioned above, SPLs in the plasma membrane have varying affinities for other lipids resulting in the generation of sub-microscopic microdomains due to non-random distribution of these compounds (Cremesti et al. 2002). While SM binds tightly with cholesterol, ceramide has low affinity for this sterol resulting in segregation and formation of exclusive ceramide-enriched microdomains (Kolesnick et al. 2000). Co-existence of microdomains may also arise as a result of ceramide mixing poorly with phospholipids. Ceramide-enriched microdomains facilitate the lamellar-hexagonal transition of lipids, which destabilize the plasma membrane lipid bilayer resulting in efflux, fusion or budding of vesicles (Cremesti et al. 2002; Blitterswijk et al. 2003).

Vesicles in the lipid bilayer are formed due to a negative curvature induced by ceramide’s small polar head group, whereas its extensive hydrogen-bonding may result in the pore formation (Siskind and Colombini 2000). Synthesis of ceramide by aSMase in the plasma membrane microdomains causes this inward vesiculation and subsequent endocytic vesicular translocation towards the mitochondria or Golgi (Blitterswijk et al. 2003). Various subsequent effects have been postulated in response to the above mentioned local changes in the plasma membrane as a result of ceramide formation, such as induction of abnormal ion fluxes, triggering conformational changes in local enzymes or receptors, alterations in transbilayer lipid movements, and enhanced movement of proteins into or from microdomains. All of these result in the activation of specific local signaling cascades which execute the biological effects of ceramide (Cremesti et al. 2002). Moreover, expression of a wide range of cell signaling proteins in membrane microdomains suggests the close involvement of these domains in signal transduction (Smart et al. 1999; Simons and Toomre 2000; Pike 2003). Studies have demonstrated roles for membrane microdomains in cellular signaling, viral and toxin entry, and protein and lipid trafficking (Ikonen and Parton 2000; Cremesti et al. 2002; Nabi and Le 2003; Parton and Richards 2003; Pike 2006).

The SMase pathway has been shown to promote membrane microdomain formation, which is one of the key sites of ceramide synthesis in response to stress (Liu and

Introduction

Anderson 1995; Bilderback et al. 1999; Yasuhara et al. 1999; Zundel et al. 2000; Cremesti et al. 2001). Generation of ceramide leads to fusion of microdomains to patches, which then coalesce to form platforms where ligated-receptor clustering and activation of downstream signaling cascades occurs (Cremesti et al. 2002). Interestingly, activation of its receptors by TNF- expels nSMase from plasma membrane microdomains, suggesting a predominant role of aSMase in regulating SM hydrolysis and ceramide generation following ligand-induced intramembrane clustering of TNF receptors (Veldman et al. 2001).

Recent studies have shown transbilayer movement of ceramide synthesized on the extracellular leaflet of the plasma membrane following exogenous addition of SM to the inner leaflet where it is phosphorylated by CERK to produce C1P (Figure 1.4). This form of ceramide production seems to be microdomain-independent, and it is possible that ceramide translocation may disrupt lipid asymmetry leading to ceramide- mediated functions such as apoptosis and/or cell cycle arrest. Furthermore, these findings imply this translocation may be spontaneous and independent of transporter proteins, and may shed light on an intrinsic property of ceramide signaling (Mitsutake and Igarashi 2007).

Figure 1.4. Transbilayer movement of ceramide. Mitsutake et al. demonstrated translocation of ceramide across the plasma membrane using a SMase/CERK-mediated system as illustrated above. (A) Exogenously added SMase hydrolyzes SM and produces ceramide, which then (B) moves across the bilayer of the plasma membrane and is phosphorylated by CERK (C) in the intracellular compartment (Mitsutake and Igarashi 2007).

Introduction

iii) Ceramide as a second messenger

In addition to its metabolic and structural role, ceramide is an important second messenger in the regulation of a number of cellular processes, including apoptosis, proliferation (Figure 1.6) (Wang et al. 1999; Yan and Polk 2001), differentiation (Okazaki et al. 1989; Dobrowsky et al. 1994), necrosis, growth arrest (Venable et al. 1995) and axonal outgrowth (Brann et al. 2002). The diversity of metabolic pathways that produce ceramide is indicative of the versatility of effector responses that can be generated by this potent mediator (Ruvolo 2001).

Ceramide has been most extensively studied for its role in triggering apoptosis following a myriad of cellular stresses. Several inducers of apoptosis, such as Fas ligand, TNF-α and chemotherapeutic agents, exert their effects by triggering pathways that lead to generation of ceramide (Futerman and Hannun 2004). Central regulators of apoptosis, such as the tumour suppressor p53, have been associated with the synthesis of ceramide in response to certain agonists (Dbaibo et al. 1998; Sawada et al. 2001; Yang and Duerksen-Hughes 2001). Key downstream signaling targets activated by ceramide include stress-activated protein kinases (SAPK), atypical protein kinase C-ζ (PKCζ), c-Jun N-terminal kinase (JNK), kinase suppressor of RAS (KSR), proteases (cathepsin D) and phosphatases (PP1 and PP2A) (Pettus et al. 2002). The protein phosphatases can, in turn, inhibit MAPK and classical protein kinases (PKA & PKB) exerting anti-growth/pro-apoptotic effects (Raines et al. 1993; Ruvolo 2001). However, the precise site where ceramide binds to these proteins awaits elucidation.

The SMase and de novo pathways are both activated in response to stress inducers, such as UV radiation, serum withdrawal, ischemia, and cytokines (Tepper et al. 1995; Liu et al. 1998). Although alkaline SMase is mainly responsible for digestion of dietary SM in the intestine, recent reports have described a role for this enzyme in inhibiting proliferation independent of apoptosis in colon cancer cells (Duan and Nilsson 2009). Conversely, aSMase and nSMase are well known to be involved in signal transduction in mammalian cells. It is of interest to note that while dihydroceramide is a direct precursor of ceramide in the de novo pathway, it does not

Introduction induce any apoptotic effects, underscoring the relevance of ceramide production (by dihydroceramide desaturase) in triggering apoptosis signaling (Pettus et al. 2002). Ceramide is also involved in cell cycle arrest in some cell lines via dephosphorylation of a key regulator of cell cycle arrest (Rb) by ceramide-activated protein phosphatases (Alberts et al. 1993; Dbaibo et al. 1995; Kim et al. 2000).

Pro-differentiation functions of ceramide and its synthesizing enzymes aSMase and nSMase have been observed in epidermal cells, whereas in myoblasts, endogenous ceramide and SMase mediate a negative feedback mechanism which limits myogenic differentiation as witnessed by downregulation of expression of myogenic differentiation markers and cell fusion rate (Mebarek et al. 2007). With regards to ceramide metabolism, ceramidases have become of particular interest because of their involvement in various genetic disorders. Four of the five major ceramidase isoforms (acid ceramidase, alkaline ceramidase-1 & -2, phyto ceramidase and neutral ceramidase) have shown altered expression levels during differentiation of keratinocytes, the exception being alkaline ceramidase-2 (Houben et al. 2007). An increase in expression and activity of alkaline ceramidase-1 and acid ceramidase has been observed in differentiating keratinocytes, while phytoceramidase and neutral ceramidase expression and activity was found to be maximal in undifferentiated keratinocytes. These data imply an essential role of ceramidases in cell differentiation and consequently maintenance of a functional skin barrier (Houben et al. 2007).

As is the case with most SPLs, ceramidase functions are tissue-specific and defined by subcellular localization. While alkaline ceramidase 1 and 2 mainly localize in the Golgi and ER fractions respectively, acid ceramidase is found within lysosomal compartments, whereas neutral ceramidase is detected in plasma membrane microdomains and the mitochondria (Ferlinz et al. 2001; Mao et al. 2001; Mitsutake et al. 2001; Mao et al. 2003; Romiti et al. 2003; Hwang et al. 2005). Generally, localization of ceramidase correlates with sites of ceramide synthesis thereby restricting potential deleterious effects of ceramide (Houben et al. 2006). Overexpression of ceramidases, in particular the acidic isoform, has been closely linked to the outcome and progression of cancer and the response of tumors to therapy (Elojeimy et al. 2007; Saad et al. 2007; Duan and Nilsson 2009). This appears to be due to reduced ceramide levels and enhanced accumulation of S1P, causing resistance to cell death and increased cell proliferation. However, ceramide is not always pro-

Introduction apoptotic or anti-proliferative; for example, ceramide has been shown to prevent neurons from undergoing programmed cell death (Song and Posse de Chaves 2003; Plummer et al. 2005).

Taken together, the body of literature indicates that changes in intracellular ceramide levels and levels of its metabolites occurs in response to agonists, which activate multiple different enzymes that vary in terms of the timing of expression and site of activity (Bourteele et al. 1998). In addition, extracellular agonists induce differential lipid accumulation with subsequent downstream effects (Nikolova-Karakashian et al. 1997). The net outcome of such stimulation depends not only on the activation of ceramide synthesis pathways (SMase, salvage and de novo), but also on the relative activities of ceramide metabolizing enzymes and alterations in other metabolically- linked bioactive lipids that may oppose actions of ceramide (for example S1P and dihydroceramide), highlighting the complexity of the SPL-ceramide metabolic network (Jarvis et al. 1994; Cuvillier et al. 1996; Auge et al. 1999). The dichotomy in structure and function between the intermediates of de novo synthesis and those of the catabolic pathway emphasizes an important aspect of SPL metabolism with regard to its role in signal transduction (Merrill 1991).

1.2.2. Sphingosine (Sph)

The importance of the salvage pathway has become clearer as more complex mechanisms of ceramide accumulation are uncovered. This pathway involves the catabolism of complex SPLs eventually breaking down into Sph in the lysosomes, which is then reused via reacylation to synthesize ceramide through the actions of acid ceramidase (Kitatani et al. 2008). Lysosomally-synthesized Sph later translocates via vesicular/non-vesicular transport to the plasma membrane and/or exits into the extracellular milieu where it becomes available for further ceramide synthesis and/or conversion to S1P (Herget et al. 2000). On the other hand, SPLs generated from the salvage pathway by reacylation of Sph are distributed to plasma membranes and subcellular organelles, which then undergo turnover with degradation and regeneration. This pathway contributes 50 to 90% of total SPL biosynthesis,

Introduction underscoring its relevance to SPL turnover/biosynthesis and cellular signal transduction (Gillard et al. 1998).

Studies have shown that this pathway is stimulated by upstream activation of PKC family members (Becker et al. 2005). It should be noted that Sph is strictly a product of SPL breakdown, contrary to its counterpart dihydrosphingosine that is mostly generated by de novo SPL biosynthesis (Kitatani et al. 2008). Sph cannot be synthesized by desaturation of dihydrosphingosine, thereby allowing the concentrations of these two long chain sphingoid bases to be differentially regulated (Rother et al. 1992). Phosphorylation of these compounds yields S1P and its dihydro form, both of which exhibit biological effects that are usually, but not always, divergent, consistent with a functional dichotomy between the dihydroSPLs of long chain bases and their reduced counterparts (Van Brocklyn et al. 1998). Interestingly, formation of Sph is not always associated with the salvage pathway and ceramide synthesis; Sph generation relies primarily on ceramidases, thus ceramidases are responsible not only for ceramide metabolism, but also the synthesis of Sph and subsequently S1P in cells (Mao and Obeid 2008).

Studies have reported induction of cell death by Sph via pathways both similar and dissimilar to those employed by ceramide (Taha et al. 2006). Exposure of cells to an apoptotic stimuli leads to an increase in ceramide levels, either via its de novo pathway or by enhanced levels of SMase activity, which results in an increase in Sph levels by ceramidase (Suzuki et al. 2004). Several enzymes are regulated by alterations in ceramidase activity and these changes are brought about by other bioactive molecules (Gangoiti et al. 2010). As discussed above, the roles of ceramidases (in particular acid ceramidase) in normal cell physiology and pathophysiology have been well documented. Acid ceramidase has been linked with Farbier disease - a rare inherited lysosomal storage disorder - atopic dermatitis, Alzheimer’s disease, cystic fibrosis, chronic hypoxia, myocardial ischemia- reperfusion, diabetic nephropathy, and in the mitogenic effect of oxidized low density lipoprotein (LDL) (He et al.; Auge et al. 1999; Bar et al. 2001; Zhang et al. 2001; El Alwani et al. 2005; Geoffroy et al. 2005; Grassme et al. 2008; Jiang et al. 2009). Acid ceramidase via its pro-proliferation/anti-cell death role has important implications for the outcome and progression of cancer (Elojeimy et al. 2007). Ceramidase inhibitors are being developed and tested for use in cancer chemotherapy and as a treatment for

Introduction other diseases such as atopic dermatitis and cardiovascular disease caused by oxLDL (Auge et al. 1999; Gangoiti et al. 2010).

Sph has been shown to induce apoptosis in a number of cells via multiple mechanisms operating in concert (Figure 1.6) (Suzuki et al. 2004). Elevated Sph levels triggers multiple signaling pathways, via activation of the caspase cascade and the release of the C-terminal-half kinase domain (KD) of PKCδ, known as Sph-dependent protein kinase (SDK) (Megidish et al. 1995; Megidish et al. 1999). Inhibition of the Akt pathway, PKC, Mg2+ -dependent and independent phosphatidate phosphohydrolase have been reported in response to Sph (Jamal et al. 1991; Gomez-Munoz et al. 1992; Smith et al. 1997). Furthermore, Sph causes activation of phospholipase D (PLD) and diacyglycerol kinase resulting in reduced diacylglycerol (DAG) and elevated phosphatidic acid (PA) levels (Sakane et al. 1989; Yamada et al. 1993; Natarajan et al. 1994).

Interestingly, exogenous treatment of Sph in multidrug resistance (MDR) leukemia cells has demonstrated anti-tumor growth properties implying potential use in cancer therapy (Klostergaard et al. 1998). Sph induces apoptosis in human epidermoid carcinoma cells expressing MDR1 transporter, despite its initial metabolites ceramide and S1P failing to do so under similar conditions, providing promising strategies and approaches to treat multidrug resistant cancers (Shirahama et al. 1997). Unlike other SPLs such as GluCer and SM, Sph does not contribute towards multidrug resistant (Klostergaard et al. 1998). Several studies have shown that SPLs exert their apoptotic effects via the caspase cascade, with early apoptotic processes triggering synthesis of some of these bioactive lipids (Sweeney et al. 1998; Pettus et al. 2002). Employing various caspase inhibitors, it has been shown that Sph acts upstream of the early caspases, whereas ceramide functions downstream of the initiator caspases but upstream from the executor caspases (Sweeney et al. 1998). Ceramide analogues and Sph, in spite of being catabolites of each other, act independently during induction of apoptosis. A role of Sph has also been reported in differentiation studies in leukemia cells upon treatment with a phorbol ester (Shirahama et al. 1997). Despite the marked differences between the mechanism of actions of Sph and S1P, Sph usually mediates most of its actions through its conversion to S1P (Desai et al. 1992; Meacci et al. 2004).

Introduction

1.2.3. Sphingosine-1-phosphate (S1P)

The key enzymes that mediate synthesis and breakdown of S1P include SPHK (which exists as two isoforms, SPHK1 and SPHK2), S1P phosphatase (S1PP, which also has two isoforms, S1PP1 and S1PP2) and S1P lyase. SPHK and S1PP are involved in synthesis and conversion of S1P from Sph (and vice versa), respectively, whereas S1P lyase regulates irreversible breakdown of S1P to palmitaldehyde and ethanolamine phosphate, providing an exit portal from the SPL metabolic pathway (Van Veldhoven 2000; Mandala 2001; Maceyka et al. 2002). S1P can also be dephosphorylated by a plasma membrane-bound Mg2+-independent and N-ethylmaleimide insensitive phosphatide phosphohydrolase, now known as lipid phosphate phosphatase (LPP) (Waggoner et al. 1996; Brindley and Waggoner 1998). Both S1PP isoforms and S1P lyase are localized to the ER, where the majority of the enzymes involved in SPL metabolism are expressed. Notably, S1P lyase and S1PP have their active sites facing the cytosolic compartment and the lumen, respectively, allowing them to access different pools of S1P.

Alternatively, it may be possible that intracellular transmembrane movement of S1P pools exists enabling both these enzymes to access the same exchangeable S1P pool (Pyne et al. 2009). In regards to SPHK, the two isoforms, SPHK1 and SPHK2, display differential spatial and functional properties with SPHK1 exhibiting a more selective substrate selection. Studies have shown that SPHK1 exists in two functional states: one is an extrinsic or agonist-induced activity involving serine 225 phosphorylation and translocation to plasma membrane, and the other an intrinsic or basal catalytic activation independent of posttranslational modifications (Pitson et al. 2003; Pitson et al. 2005). Secreted SPHK1 contributes towards generation of plasma S1P levels from circulating substrates (Liu et al. 2000; Schulz et al. 2006).

Models used to translocate SPHK1 to various subcellular localizations have shown that the location of SPHK1 rather than its catalytic activity per se may be more important in defining its function. For example, in the plasma membrane it displays oncogenic properties, whereas when redistributed to the ER or nucleus it results in either apoptosis of inhibition of DNA synthesis (Maceyka et al. 2005; Safadi-

Introduction

Chamberlain et al. 2005). This would facilitate generation of discrete compartmentalized pools of S1P with different functions in the vicinity of relevant effectors (Wattenberg et al. 2006). SPHK2, on the other hand, is mainly contained intracellularly, but also exhibits different localizations depending on cell type, varying from nucleus, cytoplasm, cytoplasmic vesicle to ER (Venkataraman et al. 2006; Alemany et al. 2007). Interestingly, some studies show localization of SPHK isoforms confined to the membrane, which is consistent with the knowledge that approximately 70% of Sph is membrane bound with only 30% being soluble at physiological pH. These enzymes, therefore, mostly metabolize Sph within the membrane compartment (Wattenberg et al. 2006; Hannun and Obeid 2008).

S1P plays a significant role in several cellular biological effects, often antagonistic to those of its direct precursors (Taha et al. 2006). Besides being a major promoter of cell survival, this blood-borne SPL is also involved in inflammation, tumorigenesis, cell growth, migration, angiogenesis, and differentiation (Spiegel and Milstien 2000; Spiegel and Milstien 2003). S1P regulates many of its biological processes as both an extracellular first messenger and an intracellular second messenger (Figure 1.5) (Payne et al. 2002). However, whether it signals inside-to-outside or outside-to-inside remains elusive, although most investigators have favored the former concept (Payne et al. 2002).

Mechanisms involved with S1P secretion are poorly understood to date, although studies have shown that the translocation of SPHK1 to the plasma membrane may facilitate the inside-to-outside signaling of S1P. There are two pathways known to generate extracellular S1P: phosphorylation of extracellular Sph by SPHK1 released from endothelial cells and release of S1P from cells (Tani et al. 2007). There are also two ATP-binding cassette (ABC) transporters associated with S1P translocation/efflux, which is discussed in further detail in Section 1.3.1 of this thesis. In this scenario, S1P would be generated by extracellular stimuli, released or exported from the cell for binding to S1P receptors and partitioning into microdomains on the outer leaflet of the plasma membrane (Spiegel and Milstien 2003). It is of interest to note that S1P can exist in a monomer–micelle equilibrium in water, and as a result it is the only signaling SPL molecule that can be found dissociated from the cell membranes in the cytosol (Garcia-Pacios et al. 2009).

Introduction

Extracellularly, S1P is a potent mitogen and component of high density lipoprotein

(HDL) that binds to a family of five specific G protein-coupled receptors (S1P1-S1P5), originally identified as part of the endothelial differentiation genes (EDG) family, differentially expressed in different tissues (Spiegel and Milstien 2000). These receptors couple to a variety of G proteins that regulate various signal transduction pathways, thus allowing S1P to stimulate diverse signaling pathways within a cell and/or in different cell types (Figure 1.5). This explains the wide range of biological outcomes regulated by S1P, with the determining factors of the outcome being the cell type, G proteins present and expression pattern of S1P receptors (Payne et al. 2002).

For example, binding of S1P to S1P1 receptor triggers cell migration and angiogenesis in various cell types (Wang et al. 1999; Yanai et al. 2000; Hobson et al. 2001;

Boguslawski et al. 2002; Idzko et al. 2002), whereas binding to S1P3 enhances survival by suppressing expression of downstream apoptotic mediators (Banno et al. 2001). Studies have also reported regulation of cell migration and membrane cytoskeleton scrambling associated with cell motility via ligand activation of S1P3.

S1P2, on the other hand, has been shown to inhibit cell migration upon activation (Okamoto et al. 2000).

A primary role of S1P2, with S1P3 playing a secondary role, has been shown in adipose tissue-derived mesenchymal cell differentiation stimulated by S1P (Nincheri et al. 2009). S1P dose-dependently stimulated differentiation of adipose tissue-derived mesenchymal stem cells towards smooth muscle cells. Indeed, S1P not only induced the expression of smooth muscle cell-specific proteins such as alpha-smooth muscle actin and transgelin, but also profoundly affected adipose tissue-derived mesenchymal stem cells morphology by enhancing cytoskeletal F-actin assembly, which incorporated alpha-smooth muscle actin (Nincheri et al. 2009). The turnover time for S1P is very rapid with a 15 minute half life, indicative of highly active synthesis and degradation pathways of S1P (Venkataraman et al. 2008). Recent studies have shown a reduction in extracellular S1P levels following its dephosphorylation by LPPs. The resulting Sph is taken up by cells then rephosphorylated via SPHK1, thereby enhancing intracellular S1P levels (Zhao et al. 2007). Although LPP has been proposed to act as an ecto-enzyme on exogenous S1P, the possibility that it may desphosphorylate intracellular lipid phosphates and regulating intracellular signaling cannot be ruled out (Brindley and Waggoner 1998).

Introduction

S1P also acts as a key bioactive molecule via its functions as a second messenger (Payne et al. 2002). It is noteworthy that cellular levels of Cer, Sph and S1P differ significantly, with highest levels exhibited by ceramide and S1P presenting the lowest levels. Thus, slight changes in ceramide levels cause considerable alterations in levels of Sph and S1P (Bartke and Hannun 2009). This makes studying S1P regulation rather challenging due to its rapid turnover rate and dual functions both as an intracellular second messenger and an extracellular ligand for a family of five G-protein-coupled receptors, three of which were recently found to be expressed in the placenta

(Johnstone et al. 2005; Hemmings et al. 2006; Hong et al. 2008). Receptors S1P4 and

S1P5 are limited mainly to cells of hematopoietic origin (Lee and Lynch 2005).

Even as a second messenger, several studies have implicated S1P in the regulation of cell proliferation, survival and inhibition of apoptosis by activating/inhibiting downstream signaling pathways involved in regulating apoptosis (Spiegel and Milstien 2000). In addition, a number of widely studied growth and development regulators, such as EGF, platelet-derived growth factor (PDGF), TNF-α, nerve growth factor (NGF), TGF-β and oestrogen have been shown to activate SPHK resulting in enhanced levels of cellular S1P (Olivera and Spiegel 1993; Meyer zu Heringdorf et al. 1999; Xia et al. 1999; Sukocheva et al. 2003; Yamanaka et al. 2004). Acute regulation of SPHK1/2 activity can be via phosphorylation, Ca2+ levels and protein- protein/phospholipid interactions resulting in an altered subcellular distribution of these isoforms (Pyne et al. 2009). ER-localized SPHK2, together with S1PP, has been shown to promote generation of ceramide via the Sph salvage pathway, whereas SPHK1-synthesized cytosolic S1P reportedly inhibits de novo ceramide production and stimulates cell proliferation (Maceyka et al. 2005). Accordingly, expression of SPHK1 is upregulated in tumor cells where inhibition of apoptosis and stimulation of invasion and proliferation is regulated by S1P (Pilorget et al. 2007).

A number of studies have indicated a role for SPHK1 in the acquisition of MDR phenotype, specifically the upregulation of the expression and function of the MDR1 (p-glycoprotein) transporter (Banno et al. 2001; Akao et al. 2006; Pilorget et al. 2007). Furthermore, over-expression of SPHK1 has been observed in a variety of tumors causing resistance to apoptosis (Minhajuddin et al. 2009). Therefore, inhibition of the anti-apoptotic SPHK1-S1P pathway is being targeted pharmacologically to enhance the efficacy of anti-cancer agents (Gangoiti et al. 2010). Studies have shown that

Introduction

SPHK inhibitors may also be important therapeutic candidates for treatment of thrombosis, atherosclerosis or hypertension, and organ inflammatory injury after shock (Lee et al. 2004; Daum et al. 2009). Moreover, since the upregulation of MDR1 expression has been associated with cell differentiation, it is likely that overexpression of SPHK1 in tumor cells leads to differentiation. S1P antibodies have recently been designed that act as molecular sponges to neutralize dysregulated S1P in relevant tissues; these have been evaluated as potential therapeutic agents in cancer and age- related macular degeneration (Sabbadini 2011). Conversely, S1P lyase has been associated with a pro-apoptotic role in response to stress inducers via an upregulation in de novo ceramide accumulation as a result of S1P depletion (Reiss et al. 2004; Saba and Hla 2004). Hence, a role in cancer cell survival may be implied during S1P lyase deficiency.

Figure 1.5. Extracellular and intracellular signaling pathways and functions mediated by S1P

(Payne et al. 2002)

Introduction

Phosphorylation of SPHK1 by recombinant ERK2 has been shown to result in its translocation to the plasma membrane. Although SPHK1 contains four putative protein kinase C phosphorylation sites, it is likely its translocation is via an indirect mechanism since purified protein kinase C reportedly has no effect on SPHK activity in vitro (Pitson et al. 2003; Wattenberg et al. 2006). ERK2-stimulated phosphorylation of SPHK1 and recruitment to membranes is regulated by association with calcium- calmodulin and phospholipids such as phosphotidylserine and phosphatidic acid (which also regulates redistribution of SPHK1 to the Golgi apparatus) (Alemany et al. 2007). Furthermore, studies suggest that PLD (which catalyses formation of phosphatidic acid) may also be an important regulator of S1P formation and signaling by redirecting SPHK1 to the Golgi apparatus (Pyne et al. 2009). In contrast, ectopically expressed LPP2 inhibits this PLD-induced localization of SPHK1 to the Golgi apparatus either by preventing translocation of SPHK1 to phosphatidic acid- derived PLD at the Golgi apparatus, or by reducing phosphatidic acid in the vicinity of PLD, thereby releasing SPHK1 from the ‘PLD trap’ (Long et al. 2005; Pyne et al. 2009). On the other hand, SPHK2 is phosphorylated and functions as a substrate of ERK1 (Hait et al. 2007).

When functioning extracellularly, S1P binds to its receptors and triggers activation of a wide range of downstream signaling targets including phosphatidylinositol 3-kinase (PI3K), Akt, ERK, p38 mitogen-activated protein kinases, small GTPases Rac and Rho, PLD and intracellular calcium mobilization resulting in the numerous biological activities that it regulates (Van Brocklyn et al. 1998; Gonda et al. 1999; Lee et al. 1999; Malek et al. 2001). One of the more widely studied functions of extracellular S1P is the regulation of cell migration and its role in angiogenesis (Payne et al. 2002). Conversely, as a second messenger the intracellular targets of S1P remain much more elusive. Studies have shown S1P activating ERK while inhibiting stress-activated kinase JNK, in line with its pro-growth anti-apoptotic role (Goodemote et al. 1995; Cuvillier et al. 1996). Furthermore, the fate of cells has been shown to depend on the dynamic balance of the opposing effects of S1P and ceramide in regulating cell growth and cell death, respectively (Mandala et al. 2000). The opposing effects of these direct SPL metabolites may be due to multiple stimuli activating precursors either directly or indirectly, the complex relationship between different lipids generated, and/or whether both substrate and product are signaling lipids (Futerman

Introduction and Hannun 2004). Investigating the enzymes involved in ceramide-S1P metabolism may, therefore, provide potential targets for development of new anticancer drugs (Cuvillier 2007). Besides growth and survival, studies have connected the SPL rheostat to several other biological processes, including calcium homestasis and allergic responses. A similar tightly regulated homeostatic balance is witnessed with ceramide-dihydroceramide metabolism and the Sph-S1P pathway related functions (Prieschl et al. 1999).

Figure 1.6. Signaling pathways and subsequent cellular responses activated by ceramide, Sph and S1P. CDases, ceramidases; MOMP, mitochondrial outer membrane permeabilization; LMP, lysosomal membrane permeabilization; FGA, fragmentation of the Golgi apparatus; S1PR, S1P receptor; SDK, SPH-dependent kinase; PP1, protein phosphatase 1; PP2A, protein phosphatase 2A; PKC, protein kinase C; SRC, Src kinase; ERK, extracellular signal-regulated kinase; AKT, protein kinase B (Mao and Obeid 2008).

1.2.4. Regulation of ABC transporter function by SPLs

Membrane lipids are involved with yet another crucial role, and that involves regulating the functional activity and/or expression of membrane proteins, in

Introduction particular ABC transporters, as they are an integral part of the plasma membrane (Van Helvoort et al. 1996; Kok et al. 2000). The lipid environment may affect membrane proteins in several ways: by determining the concentration of various exogenous/endogenous substrates and controlling their transport rates; by determining suitability of endogenous lipids as substrates; by affecting their catalytic activity; and by providing a platform for substrate loading (Bacso et al. 2004). A considerable amount of evidence suggests that components of the SPL biosynthetic/metabolic pathways are regulators of ABC protein functions. Veldman and colleagues demonstrated inhibition of MDR1 transporter activity by short chain SM, GluCer and galactosylceramide, while the opposite (stimulation) was observed in response to Sph in MDR expressing ovarian carcinoma cells (Veldman et al. 1999). However, since Sph inhibits PKC activity, and MDR1 transporter activity is affected by PKC- mediated phosphorylation, it is possible that the stimulatory effects of Sph on MDR1 activity were indirect (Sachs et al. 1996). Nevertheless, trafficking of MDR1 to the canicular plasma membrane and induction of MDR1 expression has been shown to be directly dependent on GluCer (Wojtal et al. 2006; Gouaze-Andersson et al. 2007).

The effects of exogenous SPLs on MDR1 activity do not appear to be due to non- specific membrane perturbations, since other short chain lipids fail to have any effect on the transporter. It is likely that MDR1 activity is modulated by these specific SPLs through interactions with domains of the proteins in the outer leaflet of the plasma membrane (Veldman et al. 1999). Furthermore, Pilorget et al. described the regulation of MDR1 function by SPHK and S1P in endothelial cells of the blood brain barrier (Pilorget et al. 2007). From their MDR1 substrate efflux studies they concluded that SPHK increases MDR1 expression through the generation of extra- and intra-cellular S1P. Interestingly, although S1P mediates the stimulation of MDR1 activity, on its own S1P had no effect on the expression of MDR1 (Pilorget et al. 2007). Collectively, these findings highlight a close relationship between SPLs and membrane ABC transporters, and add yet another dimension to the mechanisms regulating their biological effects in addition to aforementioned functional and metabolic roles of SPLs.

Introduction

1.3. ABC transporters

The mammalian ABC superfamily consists of 49 individual transport proteins belonging to 7 sub-families (ABCA to ABCG) involved in translocating a wide range of substrates across cellular membranes (Higgins 1992; Childs and Ling 1994; Dean and Allikmets 1995; Dean et al. 2001). While ABC proteins are typically localized in the plasma membrane, some are also expressed in intracellular membranes of the Golgi apparatus, mitochondria and ER (Dean et al. 2001; Rajagopal and Simon 2003; Ifergan et al. 2005; Solazzo et al. 2006; Tsuchida et al. 2008). ABC transporters mediate ATP-driven transmembrane efflux against a concentration gradient of a wide variety of amphiphilic ligands, including amino acids, peptides, nucleotides, glycolipids, phospholipids, sterols, polysaccharides, inorganic ions, and xenobiotics (Dean et al. 2001; Schinkel and Jonker 2003; Briggs et al. 2005). As such, they participate in diverse biological processes such as waste disposal and detoxification, cell signaling, lipid trafficking, membrane homeostasis, drug resistance and stem cell development (Jones and George 2004; Linton 2007).

Historically, research on ABC transporters focused originally on their role in xenobiotic transport (phase III metabolism), specifically their ability to confer drug resistance in tumours and influence the pharmacokinetics of a large number of pharmaceuticals (Schinkel and Jonker 2003; Szakacs et al. 2004). The major ABC xenobiotic transporters are ABCB1 (multidrug resistance protein 1; MDR1), ABCG2 (breast cancer resistance protein; BCRP) and ABCC1-3 (multidrug resistance associated protein [MRP] 1-3), all of which have been shown to limit the accumulation of cytotoxic compounds in tumour cells (Juliano and Ling 1976; Mirski et al. 1987; Cole et al. 1992; Kool et al. 1997; Veldman et al. 1999; Gottesman 2002; Doyle and Ross 2003) as well as healthy tissues (Thiebaut et al. 1987; Ayrton and Morgan 2001; Leslie et al. 2005).

Introduction

1.3.1. Endogenous SPL substrates of ABC transporters i) Mechanism of lipid efflux

Independent of their drug trafficking functions, ABC proteins are also involved in transport and regulation of a variety of endogenous compounds such as nucleotides, bile acid, porphyrins, steroids/steroid conjugates and phospho-/glyco-/sphingolipids (Figure 1.7) (Litman et al. 2001; Bodo et al. 2003; Kruh and Belinsky 2003; Schinkel and Jonker 2003). ABC proteins actively participate in maintaining lipid asymmetry across cellular membranes, including the plasma membrane (Sietsma et al. 2001). In theory, all ABC transporters that are involved in trafficking of hydrophobic drugs have the ability to transport analogs of membrane lipids (Borst et al. 2000). Following the identification of a role for the archetypal drug transporters MDR1 and MDR3 in phospholipid efflux (Smit et al. 1993; Van Helvoort et al. 1996), many of the aforementioned ABC drug transporters have also been implicated in lipid transport (Bevers et al. 1999; Borst et al. 2000; Woehlecke et al. 2003). In addition, many of the more recently identified members, particularly from the ABCA and ABCG subfamilies, have been identified as key molecules in the regulation of cellular lipid transport and whole body lipid homeostasis (Aye et al. 2009). What is more, evidence now suggests that the function of many of these proteins is, in turn, dependent on the membrane lipid milieu in which they reside.

The lipid bilayer of the plasma membrane is made up of asymmetrically arranged lipid species. While phosphatidylcholine (PC) and SM predominate in the exoplasmic (outer) bilayer leaflet, the cytoplasmic (inner) leaflet is enriched with aminophospholipids PS and phosphatidylethanolamine (PE) (Herrmann et al. 1990). This asymmetric distribution is usually preserved throughout the cellular life but in certain circumstances, such as during cell differentiation or apoptosis, asymmetry is lost through facilitated phospholipid translocation or endo-/exocytosis and related processes. Transbilayer movement of lipids is actively orchestrated by three major groups of enzymes: P-type ATPases (flippases), ABC transporters (floppases) and scramblases/lipid translocases (Bevers et al. 1999; Williamson and Schlegel 2002; Aye et al. 2009). In addition, certain regulatory molecules have been postulated to co- exist with various lipid translocases to assist with their transport functions (Ikeda et al. 2006). Consequently, disruption of the membrane equilibrium due to movement of

Introduction lipids across the membranes (lateral pressure imbalance) occurs; this is corrected when another lipid is either translocated back in the opposite direction or by expansion of one side of the membrane resulting in curvature (Sheetz and Singer 1974). The latter effect may be the underlying cause of endocytotic vesicle budding when lipids translocate towards the cytoplasmic surface (Devaux 1991; Farge et al. 1999).

ii) Efflux and trafficking of SPLs

MDR1

Although SPLs (e.g. SM, Sph, ceramide and GluCer) are localized primarily in the plasma membranes, a significant fraction can be found in the Golgi and endoplasmic reticulum. In addition to cellular efflux, trafficking of SPLs between organelles is also an important tightly regulated process. Amongst the ABCB subfamily, MDR1 is rather promiscuous in the specificity of its SPL substrates, although much of this evidence is based on transport of short chain fluorescent SPLs and may not apply to natural substrates (Neumann and van Meer 2008). Nevertheless, Eckford and colleagues presented evidence that the ability of MDR1 to transport drugs and act as a SPL floppase occur via the same mechanism (Eckford and Sharom 2005). MDR1 mediates translocation of SM and GluCer across the plasma membrane of drug resistant and normal cells (van Helvoort et al. 1997). MDR1 also mediates translocation of GluCer from the cytosolic to luminal surface of the Golgi, which is accompanied by an increase in GluCer synthase activity and synthesis of complex glycosphingolipids (De Rosa et al. 2004; Turzanski et al. 2005). Trafficking of SM and GluCer between the Golgi and plasma membrane may also be MDR1-dependent (van Helvoort et al. 1997). In cancer treatments studies using Caco-2 cells, MDR1 has been shown to prevent accumulation of cytotoxic plant and fungal sphingoid bases (except Sph) (Sugawara et al. 2004).

Introduction

MRP1

MRP1 is another ABC family member that has been reported to translocate fluorescent short-chain cholesterol and SPLs, such as SM and GluCer, similar to MDR1’s floppase mechanism. However, its transport activity is across basolateral membranes as opposed to the apical membrane efflux of MDR1 (Raggers et al. 1999). Similar to MDR1, the apparent inability of MRP1 to transport long-chain lipids throws doubt on the accuracy of the findings based on the use of short-chain analogs (Raggers et al. 1999; Raggers et al. 2000). On the other hand, SM and GluCer transport can be prevented by MDR1 and MRP1 antagonists (van Helvoort et al. 1997; Raggers et al. 1999). Furthermore, while many studies have reported MRP1- mediated transport of glutathione-conjugated substrates, others have reported no indication of any covalent glutathione-lipid conjugates amongst the short-chain SPLs transported by this protein (Raggers et al. 1999). The underlying causes of these discrepancies await further elucidation.

An important and specific role of MRP1 in S1P transport in mast cells has also been identified (Mitra et al. 2006). One of the ways S1P mediates its functions is via binding to extracellular receptors, therefore, its secretion from the cell is vital to its ability to exert its actions (Alvarez et al. 2007; Hannun and Obeid 2008; Pyne et al. 2009). Secretion of S1P in astrocytes is also inhibited by small interfering RNAs (siRNAs) specific to ABCA1, implicating ABCA1 in the efflux of this bioactive SPL (Sato et al. 2007). Whether other sphingolipids are also transported by ABCA1 remains unknown.

ABCA family

Gene knockout models have also implicated ABCA2 and ABCA3 as transporters of SM (Cheong et al. 2006; Sakai et al. 2007). ABCA2 performs a vital role in murine SM and ganglioside metabolism (Sakai et al. 2007). SM is also a substrate of ABCG1, which translocates this lipid to HDL particles (Kobayashi et al. 2006). Other ABCA subfamily members shown to mediate transport of phospholipids and GluCer include ABCA3, ABCA7 and ABCA12 (Lefevre et al. 2003; Abe-Dohmae et al. 2004; Akiyama et al. 2005). In keratinocytes, ABCA7 expression increases dramatically

Introduction with differentiation, coinciding with the accumulation of ceramide (Kielar et al. 2003). A possible involvement between ABCA7 and ceramide was also suspected when overexpression of ABCA7 in cells produced elevated ceramide levels which eventually hampered cell viability (Kielar et al. 2003). In cultures of keratinocytes from harlequin ichthyosis patients (with mutations in the ABCA12 gene), there is an abnormality in GluCer distribution which was restored by genetic correction of the ABCA12 gene (Lefevre et al. 2003; Akiyama et al. 2005). ABCA12 may, therefore, also function as a ceramide transporter, although it should be noted that a direct interaction between SPLs and ABCA7 or ABCA12 was not reported in these cases.

Figure 1.7. Overview of ABC transporters involved in lipid efflux. Schematic representation of subcellular ABC transporter localization, known acceptors and direction of transport. Black arrows represent transport direction at the plasma membrane. Vectorial transport by intracellular transporters has not been firmly established. Apo, apolipoprotein; HDL, high density lipoproteins; LDL, low density lipoproteins; Chol, cholesterol; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PS, phosphatidylserine; Cer, ceramide; N-retinyl-PE, N-rentinyl phosphatidylethanolamine; GluCer, glucosylceramide; S1P, sphingosine-1-phosphate; SM, sphingomyelin; LPA, lysophosphatidic acid (Aye et al. 2009).

Introduction

1.3.2. Effects of membrane SPLs on ABC transporter activity

In addition to regulating membrane lipid composition and distribution, the activity of ABC transporters can in turn be modulated by the lipid environment in which they reside. Studies have demonstrated roles for membrane microdomains in cellular signaling, viral and toxin entry, and protein and lipid trafficking (Ikonen and Parton 2000; Cremesti et al. 2002; Nabi and Le 2003; Parton and Richards 2003; Pike 2006). ABC transporters MDR1, MRP1, ABCA1, ABCG1 and ABCG2/BCRP are amongst the various proteins expressed in membrane microdomains and their complex interactions with membrane lipids and/or proteins determine their overall cellular effects.

i) MDR1

Expression of MDR1 has been identified in various different kinds of membrane microdomain fractions with different functions based on its localization (Bacso et al. 2004) and domain properties (Dos Santos et al. 2007). As microdomains are highly enriched in SM, ceramide and cholesterol, the activity of domain-associated proteins, including ABC transporters, is sensitive to variations in membrane content of these lipids (Bacso et al. 2004; Pike 2004). Drug binding, transport and ATPase activity of MDR1 expressed in membrane microdomains have been shown to be influenced by cholesterol (Saeki et al. 1992; Romsicki and Sharom 1999; Wang et al. 2000; Rothnie et al. 2001). While some studies reported MDR1 function to be cholesterol-dependent, others argue that MDR1 is functional in the absence of cholesterol (Romsicki and Sharom 1998; Lu et al. 2001; Bacso et al. 2004). In contrast to MDR1, MRP1 has been localized to microdomains with similar lipid/protein compositions exhibiting much less heterogeneity (Hinrichs et al. 2004; Radeva et al. 2005). Moreover, reports show that only severe cholesterol depletion (<40 %) was able to significantly reduce MRP1 activity (Klappe et al. 2004; Marbeuf-Gueye et al. 2007).

Introduction ii) ABCA1

The association of ABCA1 transporter with various types of membrane microdomains is cell type specific (Schmitz et al. 2000) (Mendez et al. 2001; Drobnik et al. 2002). Overexpression of ABCA1 has been shown to result in a marked reduction in the amount of cholesterol and SM in TRMs (Landry et al. 2006). Altering ABCA1 levels also impairs Akt signaling which is known to be regulated by microdomain composition and structure. It is interesting to note that mutations in the ATP binding domain of ABCA1 fail to induce cholesterol or SM redistribution, and do not impair Akt signaling. (Mendez et al. 2001)

iii) ABCG family

Evidence of ABCG1 expression in microdomains was identified by reports that alterated SM levels resulting in changes in ABCG1 function (Sano et al. 2007), although it should be noted that unlike ABCA1, ABCG1 has not been directly isolated from membrane microdomains. Studies have described cholesterol translocation to the extracellular leaflet of the plasma membrane by both ABCA1 and ABCG1 in macrophages in an apoA-I-dependent and/or –independent manner, a process which seems to be dependent on SM content in the plasma membrane (Wang et al. 2004; Vaughan and Oram 2005). Ceramide is also a regulator of cholesterol efflux via enhancement of plasma membrane-bound apoA-I (Witting et al. 2003). Paradoxically, while reduced SM levels enhance efflux of cholesterol by ABCA1, it may decrease ABCG1-mediated cholesterol efflux (Nagao et al. 2007). Cholesterol efflux is preferentially accompanied by flopping PC and SM by ABCA1 and ABCG1, respectively (Kobayashi et al. 2006; Nagao et al. 2007).

Storch and colleagues have shown in canine kidney epithelial cells that ABCG2/BCRP activity is regulated by the cholesterol content of membrane microdomains (Storch et al. 2007). BCRP expression was detected mainly in membrane microdomains, but interestingly, localization remained unaffected following cholesterol depletion (Dos Santos et al. 2007; Storch et al. 2007). In agreement with these findings, studies on human and insect membrane vesicles demonstrate variable BCRP stimulation depending on cholesterol loading and

Introduction depletion (Pal et al. 2007; Telbisz et al. 2007). Given the pivotal role of BCRP in the absorption, distribution and excretion of xenobiotics, recognizing the significance of cholesterol in modulating its activity could have important pharmacotherapeutic implications.

1.4 ABC transporters, SPLs and trophoblast differentiation

The placenta performs a multitude of functions, one of which is a complex transport system that is vital for normal development of the fetus and the maintenance of pregnancy (Evseenko et al. 2006). The syncytium facilitates this transport function by expressing various transporters differentially in a polarized manner in the fetal-facing basal membrane and the maternal-facing apical brush-border membrane (Ganapathy et al. 2000). During villous syncytial formation, an upregulation in expression of BCRP has been demonstrated (Evseenko et al. 2006). Silencing BCRP in trophoblast-like BeWo choriocarcinoma cells undergoing FSK-induced differentiation downregulated differentiation and fusion, with a subsequent increase in apoptosis (Evseenko et al. 2007c). Independent of xenobiotic transporter, studies have implicated a role for BCRP in the regeneration potential of cells in various organs, since silencing expression of these transporters in mice reduced their stem cells subpopulation (Jonker et al. 2005). These findings are in agreement with Evseenko et al. which implicate BCRP in the regeneration of placental sycyntial epithelium (Evseenko et al. 2007c). BCRP has also been associated with the regulation of trophoblast survival during pregnancy complications (Evseenko et al. 2007a). During hypoxia, toxic products are effluxed from the cytoplasm by BCRP, thereby conferring resistance to abnormal physiological conditions (Krishnamurthy et al. 2004).

Evseenko et al. successfully demonstrated that the expression of BCRP during differentiation was positively regulated by EGF, insulin-like growth factor II (IGF II) and estrogen, and negatively regulated by cytokines (Evseenko et al. 2006; Evseenko et al. 2007b). Regulation of BCRP expression by progesterone has been described in BeWo cells (Wang et al. 2008); however, these findings did not replicate in primary trophoblasts (Evseenko et al. 2007b). This may be due to already existing high levels

Introduction of endogenous progesterone in primary cells, resulting in receptor saturation, rendering addition of exogenous progesterone without affect.

BCRP was also shown to protect trophoblasts from apoptosis induced by TNF-α and short chain ceramides (Evseenko et al. 2007a); silencing expression of this protein enhanced accumulation of endogenous ceramides in cells undergoing cytokine- induced stress (Figure 1.8).

Figure 1.8. Effects of TNF-α/IFN-γ on C16, 18, 20 and 24 ceramide accumulation. BeWo cells were transiently transfected with Stealth scrambled siRNA or BCRP siRNA; 48 h after transfection cells were treated with TNF-α (50ng/ml)/IFN-γ (100 U/ml) for 3, 12 and 24 h in either complete media with 10% FCS (A,B), or serum-free media supplemented with 0.5% human albumin (C, D). Levels of ceramides were measured by LC-MS/MS in cells and culture media and results shown as the sum of all 4 ceramide species normalized to cholesterol levels in the same samples. A) Levels of ceramides in BeWo cells cultured in complete media. B)Levels of ceramides in conditioned media. C) Levels of ceramides in BeWo cells cultured in serum-free media supplemented with 0.5% human fatty acid-free albumin. D) Levels of ceramides in serum-free conditioned media. Cholesterol content was stable through all time points independent of TNF-α/IFN-γ treatment. To calculate ceramide levels in the media, basal concentrations of ceramides in FCS containing media (before experiments) were deducted from the total levels detected after incubation with the cells. Statistical analysis was carried out by one- way ANOVA with repeated measurements; *P >0.05 vs. control siRNA (Evseenko et al. 2007c).

Introduction

Interestingly, the protective effect of BCRP was found to be restricted to the extrinsic apoptotic pathway, since suppressing BCRP expression had no effect on apoptosis mediated by activators of the intrinsic pathway (Evseenko et al. 2007a). Since EGF has been widely studied as a survival factor in trophoblast cell cultures, particularly cells undergoing TNF-α/ceramide-induced apoptosis, it is possible that EGF and other survival factors may mediate their effect in part by enhancing expression of BCRP. Moreover, exposure to inflammatory cytokines in BCRP-silenced cells increased externalization of PS, which as explained above is seen in earlier stages of apoptosis. Taking these findings into account, and in light of the fact that BCRP possesses aminophospholipid flippase functions, it could potentially explain the increase of PS efflux in BCRP-silenced cells. This implies that the link between BCRP and trophoblast differentiation/survival may lie with an interaction between the transporter and the recovery of the plasma membrane lipid architecture following fusion (Evseenko et al. 2007a; Evseenko et al. 2007c). Interestingly, reduced levels of BCRP have been reported in pregnancies with fetal growth restriction (FGR), a condition associated with impaired placental function and increased apoptosis (Evseenko et al. 2007a). These findings may have significant clinical relevance and therapeutic potential; for example, BCRP expression may be responsive to growth factors, pregnancy-related steroids and cytokines to augment its protective capacity and improve placental health and function.

1.5 Summary, aims and hypotheses

The coordination of both lipid- and protein-mediated signals is essential for proper cell execution (Taha et al. 2006). SPLs have been widely implicated in the regulation of apoptosis and several other cellular activities, in particular differentiation. Despite the well defined role of SPLs in cell differentiation, there is remarkably little information on the role of SPL biosynthesis, metabolism and actions in trophoblast differentiation and fusion. Recent studies have shown that SPLs are substrates of ABC transporters (De Rosa et al. 2004). Evseenko et al. proposed a role of BCRP tranporter in the regulation of trophoblast differentiation by showing an alteration in expression during syncytialization (Evseenko et al. 2006). Blocking these transporters in

Introduction preliminary studies showed reduced syncytial formation, sensitization of trophoblasts to cytokine-induced apoptosis and an increase in the accumulation of ceramide.

In light of this knowledge gap, and the recent data on the association between ceramides, BRCP and trophoblast differentiation, the aims of this thesis were to explore the relationship between placental BCRP transporters and cellular SPL levels during differentiation of trophoblasts, and the biosynthesis and roles of SPLs during this process as well. Elucidating the mechanisms that govern villous trophoblast differentiation and syncytialization, and the expression and function of SPLs and ABC transporters in this process, will improve our understanding of a key aspect of placentation and may shed light on the etiology of a range of pregnancy disorders.

This thesis is constructed around the following hypotheses:

1. Sphingolipids are involved in the autoregulation of trophoblast differentiation and syncytialization.

2. Cellular sphingolipid levels in trophoblasts alter with differentiation as a consequence of changes in abundance of enzymes involved in sphingolipid biosynthesis and metabolism.

3. BCRP (ABCG2) transporter, via its effects on distribution of apoptotic and mitogenic sphingolipids, serves a fundamental role in modulating trophoblast and differentiation.

To explore these hypotheses, we assessed changes in various sphingolipid-related metabolic enzymes during biochemical and morphological trophoblast differentiation, using a well-validated model of in vitro differentiation, in correlation with intracellular ceramide, Sph and S1P levels. We investigated if there was an association between BCRP knockdown on trophoblast differentiation and levels of SPL biosynthesis enzymes. We also manipulated concentrations of exogenous ceramide,

Introduction

Sph and S1P and altered internal levels of these SPLs via inhibition of key metabolic enzymes to study their effects on functional and morphological differentiation.

Methods

CHAPTER 2. MATERIALS AND METHODS

2.1. Reagents

Reagents and consumables Supplier

Medium 199 with Earle’s salts and L- Mediatech Inc (Manassas, VA, USA) glutamine

Recombinant human IFN-γ, DNAse I and Roche Diagnostics (Mannheim, complete protease inhibitors, M30 Germany) antibody (used as marker of execution stages of apoptosis)

Fetal bovine serum (FBS) and fatty acid Bovogen (Victoria, Australia) free bovine serum albumin (BSA)

Monoclonal anti-β actin, peroxidase Sigma-Aldrich (St. Louis, MO, USA) conjugated goat anti-mouse antibody, bicinchoninic acid (BCA) reagent, Hoechst 33258, goat anti-rabbit peroxidase-conjugate, cholesterol, ethyl acetate, methanol, ethanol, chloroform, isopropanol, 3,3´,5,5´- tetramethylbenzidine (TMB) susbtrate, methylumbelliferyl phosphate (MUP) reagent, Sigma fastTM 3,3– diaminobenzidine (DAB), and all other cell culture grade chemicals

Insulin/transferrin/selenium supplement Invitrogen (Carlsbad, CA, USA) (100 x), human recombinant epidermal

Methods growth factor (EGF), stealth siRNA duplexes (for human glial cell missing (GCM)-1, acid sphingomyelinase (aSMase), ceramidase and sphingosine kinase (SPHK)-1), scrambled siRNA negative control, F12-K Nutrient Mixture, Kaign’s Modification (1 x), dispase-II, antibiotic/antimycotic (100 x), Dulbecco’s phosphate buffered saline (D-PBS), 0.05% trypsin-EDTA, Zymax goat anti rabbit/mouse Cy3 conjugate antibodies, Pure Link RNA mini kit, and SYBR safe DNA gel stain

Nitrocellulose Hybond membrane Amersham Pharmacia Biotech (Buckinghamshire, UK)

Rabbit anti-ceramide kinase (CERK), Abcam (Cambridge, MA, USA) rabbit anti-E-cadherin (HECD-1), rabbit anti- SPHK1 & SPHK2, antiS1PP2 and mouse anti-hCG.

N-palmitoyl-D-erythro-sphingosine (C16 Avanti Polar Lipids (Alabaster, ceramide), N-octanoyl-D- AL,USA) erythrosphingosine (C8 ceramide) and N- palmitoyl (D31)-D-erythro-sphingosine (C16-D31 ceramide), D-erythro- sphingosine (synthetic), D-erythro- sphingosine (C17 base), D-erythro- sphingosine-1-phosphate (C17 base) and D-erythro-sphingosine-1-phosphate.

Methods

Rabbit anti- aSMase Exalpha Biologicals Inc., (Massachusetts, U.S.A.)

Mouse anti-ceramidase (ASAH1; clone Abnova Corporation (Taipei, Taiwan) 2C9)

Fluorescent-labelled goat anti-mouse Biolegend (San Diego, USA) DyLight 594

Forskolin, okadaic acid and tautomycin Enzo Life Sciences (New York, USA)

ProFection® Mammalian Transfection Promega (Madison, WI, USA) System, ImProm-II™ Reverse Transcription System and Go Taq hot start polymerase, Caspase-Glo ® 8 Assay and Caspase-Glo® 3/7 Assay

Mouse anti-BCRP (clone BXP 21) Chemicon (Temecula, USA)

Anti-cytokeratin-7 antibody DacoCytomation (Glostrup, Denmark)

CAY10466/B13 (N-[(1R,2R)-2-hydroxy- Cayman Chemical (Michigan, U.S.A). 1-(hydroxymethyl)-2-(4- nitrophenyl)ethyl]-tetradecanamide),

Fuminosin B1 2-(p-Hydroxyanilino)-4-(p- chlorophenyl) thiazole

Percoll GE Healthcare (Wisconsin, U.S.A.)

Fumitremorgin C, Dimethyl-D-erythro- Calbiochem, EMD Biosciences Inc.

Methods sphingosine, ERK1/2 inhibitor PD98059 (Darmstadt, Germany) (2′-Amino-3′-methoxyflavone), p38MAPK inhibitor SB203580 [4-(4- Fluorophenyl)-2-(4- methylsulfinylphenyl)-5-(4-pyridyl)1H- imidazole], JNK inhibitor SP600125 (anthra[1,9-cd]pyrazol-6(2H)-one), PKA inhibitor H89 (5- Isoquinolinesulfonamide), SPHK1 inhibitor 2-(p-Hydroxyanilino)-4-(p- chlorophenyl) thiazole, PI3K inhibitor LY294002 (2-(4-morpholinyl)-8-phenyl- 4H-1-benzopyran-4-one), D-erythro- MAPP (1S,2R)-D-erythro-2-(N- Myristoylamino)-1-phenyl-1-propanol dNTPs Bioline (London, UK)

EvaGreen Biotium Inc. (Hayward, CA, USA)

Analytical grade chemicals Calbiochem, EMD Biosciences Inc. (Darmstadt, Germany)

Anti-phosphoAkt (Ser473), anti-total Akt, Cell Signaling Technology, Inc. anti-phospho-SAPK/JNK (Danvers, MA, U.S.A.) (Thr183/Tyr185), and anti-total SAPK/JNK.

Table 2.1. List of Reagents

Methods

2.2. Isolation and culture of term human placental trophoblast cells

Human term placentae were obtained from King Edward Memorial Hospital from women at term undergoing elective Caesarean sections. Informed consent from the mothers and ethical approval from the institutional human ethics committee was obtained prior to collection of the placentae.

2.2.1. Buffers and Solutions

Red cell lysis buffer: 2.675 g/L 50 mM ammonium chloride, 0.84 g/L 10 mM sodium hydrogen carbonate, 0.029 g/L 0.1 mM EDTA dissolved in 250 ml MilliQ water.

HBSS buffer: 8 g/L sodium chloride, 0.4 g/L potassium chloride, 0.35 g/L sodium hydrogen carbonate, 0.06 g/L potassium hydrogen phosphate, 0.048 g/L sodium hydrogen phosphate in 1 L MilliQ water.

Dispase digest solution: 1.25 g/L in HBSS buffer.

Percoll: gradients (5%, 20%, 30%, 35%, 40%, 45%, 50% and 60%) in M199 media.

Complete Cell culture medium: M199 supplemented with 10% serum, 1% antibiotic/antimycotic, 1% insulin/transferring/selenium and 10 ng/ ml EGF.

2.2.2. Trophoblast isolation

Primary trophoblast cells were isolated from term placenta by dispase digestion as previously described (Singh et al. 2011). Square chunks of villous tissue, approximately 2 to 5 mm thick, were cut from the cotelydons of term placentae and thoroughly washed in D-PBS. While cutting out villous pieces, precautions were taken to avoid connective tissue, sinews, blood vessels and discolored areas where possible. After collecting approximately 100 ml of tissue, the tissue was digested in dispase II (0.25 % w/v in HBSS buffer) for 1 h at 37°C, with DNAse I (2 mg/ml) added 15 min prior to the end of digestion. Tissue digests were then filtered through

Methods

100 μm cell filters, centrifuged at 300 g for 10 min and erythrocytes removed by incubating cell pellets in lysis buffer. Following a quick wash step with culture media, the liberated cells were given an acute exposure to 1% trypsin-EDTA for 1 min before stopping the reaction with FBS and pelleting cells at 1000 g for 5 min. Cells were subjected to centrifugation at 1,200 g for 20 min on a discontinuous Percoll gradient and cells which migrated between 20 and 40% Percoll bands (trophoblasts) were collected and layered on a new Percoll gradient and centrifuged as before and suspended in complete M199 media. Aliquots (10 μl) of re-suspended cells were diluted 1:9 in 0.1% trypan blue solution, the exclusion of which determines viability of cells, and counted using a haemocytometer. Placental variability was accounted for by the varying cellular yields, which ranged from 50 to 150 million. Cells were plated in supplemented M199 culture media at a density of 1 million cells/ml. Following overnight incubation in an incubator equilibrated in 95% air/5% CO2, media was replaced (to remove non-adherent cells) with supplemented M199 media and differentiation experiments were carried out.

Exogenous treatments. Cells cultured in 6 well, 24 well and 96 well plates were treated with the following treatments for up to 5 days of culture (media changed daily) at 37˚C post isolation, after which cells lysates and culture media were collected for further experimental analysis. Vehicles used were culture media with upto 0.1% of EtOH or DMSO depending on respective treatments.

Treatment Concentration

C8 Ceramide 10 µM

Sphingosine 10 µM

S1P 10 µM

aSMase 0.01 IU/ml

Methods

B13 (ceramidase inhibitor) 25 µM

D-erythro-MAPP 10 µM

Fuminosin B1 10 µM sphingosine kinase inhibitor 10 µM

DMS 5 µM

SP600125 (JNK inhibitor) 10 µM

SB203580 (p38 inhibitor) 2 µM

PD98059 (ERK1/2 inhibitor) 20 µM

LY294002 (PI3K inhibitor) 20 µM

Okadaic acid (PP2A inhibitor) 1 nM

Tautomycin (PP1 inhibitor) 10 nM

Fumitremorgin C (FC) 2.5 µM

Table 2.2. Final concentrations used of various test substances

Methods

2.3. Gene Expression Analyses

2.3.1. RNA extraction

Total RNA was extracted from cells or tissues and DNAse treated using the Pure Link RNA mini kit according to the protocol outlined below. Approximately 150 to 300 μl of RNA lysis solution was added per well to 24- or 6-well plates, respectively. Equal volume of 70% ethanol in nuclease free water was then added to the lysates and the solution added to a filter cartridge and centrifuged at 14,000 rpm for 1 min, with the flow through being discarded. Approximately 700 μl of Wash solution 1 was then added to the filter cartridge and centrifuged as before. Wash solution 2 and 3 were then added and centrifuged consecutively as before. The RNA was then eluted by adding 10 μl of elution solution to the filter cartridge and the RNA collected for genomic DNA removal.

2.3.2. Genomic DNA removal using DNAse

DNAse I solution (1 μl) and DNAse I buffer (0.1 x volume of total RNA) were added to the extracted RNA and incubated for 20 min at 37°C. Reaction was terminated by adding the inactivating agent (0.1 x total RNA volume) and incubated for 2 min at room temperature. The inactivating agent was then removed by centrifugation at 14,000 g for 1 min and the purified RNA aliquoted into a new tube. The quality and concentration of the obtained RNA was measured using the Nanodrop 1000 (Thermo Fisher Scientific Inc, Waltham, MA, USA). A ratio of 1.8 – 2.0 at 260/280 nm was considered to be indicative of adequate quality.

2.3.3. Reverse transcription and cDNA synthesis

RNA was reverse transcribed using the ImProm-II™ Reverse Transcription System according to manufacturer’s instructions (Promega). Briefly, up to 1 μg of total RNA was added to a PCR tube with 0.5 μg of random primers and incubated for 5 min at 70 °C. The following were then added to each RNA sample: 3.7 μl of nuclease free water, 1:5 diluted reaction buffer, 6 mM MgCl2, dNTP mix (0.5 mM each), 20 U of

Methods recombinant RNAsin ribonuclease inhibitor and 1 μl of reverse transcriptase, and incubated for 5 min at 25 °C (annealing), 1 h at 42 °C (extending) and 15 min at 72 °C (reverse transcriptase inactivation).

2.3.4. Quantitative real-time PCR (qPCR) qPCR (see Table 2.1 for primer information) was performed for genes of interest in triplicate on 0.2 – 1 μg of cDNA using 1:5 stock of PCR buffer, 2 mM MgCl2 (2 μl/reaction), 0.1 U/μl dNTP (0.5 μl/reaction), 0.2 μM primers (0.5 μl/reaction), 1:20 Eva Green (1.25 μl/reaction) and 0.025 U/μl Go Taq polymerase (0.125 μl/reaction). PCR amplification and detection was performed on a Rotorgene 3000 (Corbett Industries, Sydney, Australia) using the initial denaturation condition of 95°C for 2 min, followed by 40 cycles at 95°C, 60°C and 72°C for 30 seconds each. Exponential amplification of all PCR reactions ranged between 1.95 – 2.00 across 7 serial log dilutions of the template. Amplification of a single product was confirmed by melting curve analysis and visualization of the product by gel electrophoresis on a 2 % agarose gel. mRNA expression was quantified using the comparative threshold cycle (Ct) method for relative quantification (2-ΔΔCt), normalized to the level of the housekeeping gene SDHA.

Gene Primers Tm Product Amplification length Efficiency

aSMase Forward: 5’-GAGAGATGAGGCGGAGACC-3’ 53.23 149 0.96

Reverse: 5’-CTTCATTGAGGGCAACCACT-3’ 52.24

Ceramidase Forward: 5’-ACGTTGGTCCTGAAGGAGG-3’ 53.11 105 1.02

Reverse: 5’-CCTTAGTCCTCCTGGCTGC-3’ 53.67

SPHK1 Forward: 5’-GTGGCTGTAGGAGCACCG-3’ 54.48 98 1.06

Methods

Reverse: 5’-CCAGGCCACTGTAGGGAAC-3’ 53.88

SDHA Forward: 5’-TGGGAACAAGAGGGCATCTG-3 53.76 86 0.98

Reverse: 5’-CCACCACTGCATCAAATTCATG-3’ 52.70

GCM1 Forward: 5’-GCATAGGAATCTGGCCACTC-3’ 52.33 136 1.01

Reverse: 5’-GGCCTGAACTTATCATGGAA-3’ 49.28

Table 2.3. Gene specific primers for quantitative PCR. Primers were designed using the Primer-BLAST program (http://www.ncbi.nlm.nih.gov/tools/primer-blast/). Primers were designed to span at least one intron on the corresponding genomic DNA and to give amplicon sizes between 50 and 200 bases.

2.3.5. Visualisation of PCR products by gel electrophoresis

2% agarose (w/v) was dissolved in Tris-Acetate-EDTA buffer (4 mM Tris, 5.7% glacial acetic acid, 1 mM EDTA) by heating in microwave at 1 min intervals. The mixture was allowed to cool prior to the addition of 0.01% SYBR safe DNA gel stain and poured into a gel tray. Once the gel was set, it was placed in an electrophoresis buffer and filled with Tris-Acetate-EDTA buffer until the gel was just submerged. PCR products and the DNA ladder were diluted with 6 x loading dye and loaded into the wells (10 µl/well). The gels were then run at 90 V for approximately 50 min and the bands visualised on an ImageQuant 350 digital imaging and quantification system (GE Healthcare).

Methods

2.4. Protein Expression Analyses

2.4.1. BCA assay i) Buffers and Solutions

RIPA buffer: 25 mM Tris-HCl (pH 7.8), 1% Triton (x100), 2 mM EDTA, Protease Inhibitor tablet, 1% SDS and 1% Deoxycholic acid.

BCA stock solution: 4 mg/ml bovine serum albumin (BSA) diluted in RIPA buffer

BCA working reagent: Copper sulphate (4% stock) diluted in BCA reagent (1:50) ii) BCA standard curve preparation and loading samples

For determination of protein concentrations in tissue/cell homogenates, a set of standard solutions were prepared by making a 1:2 serial dilution of 4 mg/ml BSA stock solution in RIPA buffer (1:1), giving a set of solutions with the following protein concentrations: 2000, 1000, 500, 250, 125, 62.5, 31.25, 0 μg/ml. 25 μl of the standards were loaded (in duplicate) into the first two columns of a 96-well microtitre plate (Greiner Bio-one, Gamburh, Germany). Lysed trophoblast cell protein samples were diluted 1:5 in RIPA buffer and 25 μl of the prepared samples were added in duplicates into subsequent columns. BCA reagent (200 μl) was added to each well, and the plate was incubated on an orbital shaker at room temperature for approximately 30 min. The plate was placed onto a Spectramax plate reader (Molecular Devices) set to read an absorbance wavelength of 562 nm. Standard curves were prepared by measuring the absorbance of BSA calibrators at 562 nm. The calibration curves were linear (r2>0.99) over the concentration range tested (0 – 2000 μg/ml) and the precision of all standards was acceptable (CV < 5 %). The protein concentration was determined using VersaMax plate reader (Molecular Devices, CA, U.S.A.).

2.4.2. Immunoblotting

Immunoblotting was performed on total cellular protein lysates. Crude membrane preparations from trophoblasts and BeWo cell lysates were prepared in RIPA buffer

Methods from cells cultured in 6 well plates daily for up to seven days in culture using a Bandelin Electronic Sonicator (Bandelin Electronic, Berlin, Germany). After protein concentrations were measured by BCA assay, proteins samples (20-40 μg, equal loadings per experiment) were separated under reducing conditions on a 4-12% BisTris precast polyacrylamide gradient gel (Invitrogen, Carsbad, USA) and transferred for 90-120 min at 30 V to nitrocellulose membrane in an XCELLTM transfer module (Invitrogen, Carsbad, USA). Membranes were blocked in 1% non-fat milk powder in phosphate buffered saline (PBS) with 0.1% Tween 20 (PBS-T) for 1 h and incubated overnight at 4ºC with monoclonal antibodies diluted as follows: anti- BCRP (1:500), -aSMase (1:200), -ceramidase (1:750), -CERK (1:500), -SPHK1 (1:200), -S1PP2 (1:400) or β-actin (1:5000). Membranes were then washed in PBS-T 3 x for 10 min and incubated with horseradish peroxidase-conjugated goat anti-mouse antibody (1:5000) diluted in 2% non-fat milk powder in PBS-T, and visualised using a sensitive enhance chemiluminescence detection kit (SuperSignal West-Femto Substrate, Pierce Chemical, Rockford, IL, USA). Resultant images were quantified on the ImageQuant 350 digital imaging and quantification system according to manufacturer’s instructions.

2.4.3. Human chorionic gonadotropin (hCG) ELISA.

Secretion and expression of hCG was determined by in-house ELISA. Briefly, culture media and lysates from cells cultured in vitro (5 x 105 cells/well in 24 well plate for trophoblasts; 3.5 x 105 cells/well for BeWo cells) were collected at different time points to determine levels of hCG secretion and expression respectively. Plates were coated with mouse anti-hCG capture antibody (1:500) in blocking buffer (0.5% BSA, 10% Newborn calf serum in 1x wash buffer) overnight at 4˚C, followed by incubation with samples and standards (0.137 mIU/ml - 100 mIU/ml) for 30 min. Exposure to detection antibody (rabbit anti-hCG; 1:5000) was carried out for an hour prior to incubation with horseradish peroxidase-conjugated antibody (1:2,500) for 30 min. After washing, plates were incubated with TMB substrate (100 µl/well) for 5-10 min at room temperature before stopping the reaction by addition of 2 N sulphuric acid (50 l/well) and absorbance was read at 450 nm on a VersaMax plate reader (Molecular Devices, CA, USA).

Methods

2.4.4. PLAP assay

To determine PLAP activity, trophoblast cell lysates collected from different time points in culture were incubated with MUP reagent (1 mg/ml) diluted 1:100 in alkaline phosphatase buffer to give a working solution of 0.01 mg/ml, as previously described (Aye et al. 2011). Samples were diluted 1:1 in alkaline phosphate buffer prior to incubation with MUP reagent for an hour at 37˚C. Level of PLAP activity (represented by fluorescence signal) was measured using a FLx800 Multi-Detection Microplate reader (BioTek® Instruments, Inc., Vermont, USA) at excitation and emission wavelengths of 360 and 460 nm, respectively.

2.5. Immunocytochemical detection

Following treatment with various SPL compounds, enzyme- and signaling pathway- inhibitors at the indicated concentrations for 24, 48 or 72 h, cells were fixed in 4% paraformaldehyde (or icecold methanol in the case of M30 immunostaining) for 10 min. Cells were incubated in blocking buffer (5% newborn calf serum/1% BSA in PBS-T) then incubated in either anti-cytokeratin-7 (trophoblast specific marker- 1:250), anti-E-cadherin (cell fusion marker -1:50), or M30 antibody (apoptosis marker - 1:100) for 1 h in blocking buffer at room temperature. Following washes in PBS-T (3 x buffer change, 5 min each) cells were incubated with fluorescent-labelled goat anti-mouse or peroxidase-labelled detection secondary antibody for 2 h at 1:400 dilution. Visualisation of the positive cells incubated with peroxidise conjugated secondary antibody was achieved using DAB peroxidase substrate. As a negative control the primary antibody was omitted. Each experiment was performed at least 3 times. Nuclei were stained using Hoechst 33258 (5 μg/ml) for 5 min and cells visualised using the Nikon Eclipse Ti inverted microscope (Nikon Imaging Inc., Tokyo, Japan).

Purity of trophoblast cultures was determined by analysing cytokeratin-7 expression, a trophoblast specific marker. Cells were incubated with anti-vimentin (1:50), a fibroblast

Methods marker, to confirm lack of contamination by other cell types. A portion of > 95 % trophoblasts in the total cell population was considered as sufficient purity.

Fusion of the mononuclear CTBs to form the syncytiotrophoblast layer was assessed by quantifying the distribution of E-cadherin and nuclei in cells after fixation and immunostaining as previously described with modifications (Orendi et al. 2010). Staining of E-cadherin, present at the intercellular boundaries in aggregated cells, progressively disappears with syncytium formation. Cell fusion event was defined as the number of cells with more than 1 nuclei within each syncytium. Analysis was carried out by counting the ratio of multinuclear cells to total number or nuclei per visual field, in 3 fields of view per well at 200 X magnification in duplicate wells.

Detection of cytokeratin 18 neo-epitope, which is cleaved by activated caspase 3, with the M30 antibody was used to determine caspase 3 activity (Black et al. 2004).

2.6. Measurement of caspase activity

2.6.1. Caspase 8

Caspase 8 activity was measured using Caspase-Glo® 8 Assay from Promega Corporation (Madison, USA) according to the manufacturer’s protocol. Cells were seeded in 96-well plates incubated with test agents. Control wells containing culture media without cells were included to obtain a value for background luminescence value. After incubation, plates were kept at room temperature for 30 min, and then 50 μl of Caspase-Glo® 8 Reagent was added to each well. After shaking for 30 seconds and incubation at room temperature for 30-60 min, luminescence was measured on a FLx800 Multi-Detection Microplate reader (BioTek® Instruments, Inc., Vermont, USA).

2.6.2. Caspase 3/7

Caspase 3/7 activity was measured using Caspase-Glo® 3/7 Assay from Promega Corporation (Madison, USA) according to the manufacturer’s protocol. Cells were seeded in 96-well plates incubated with test agents. Control wells containing culture

Methods media without cells were included to obtain a value for background luminescence value. After incubation, plates were kept at room temperature for 30 min, and then 50 μl of Caspase-Glo® 3/7 Reagent was added to each well. After shaking for 30 seconds and incubation at room temperature for 30-60 min, luminescence was measured on a FLx800 Multi-Detection Microplate reader (BioTek® Instruments, Inc., Vermont, USA).

2.7. Cell viability (MTT) assay

The 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay, based on the method of Mosmann et al. was used to assess cell viability in 96-well plates after 24, 48 and 72 h of exposure to the various test agents (Mosmann 1983). At the desired time points, cells were incubated with MTT reagent (5 mg/ml in PBS) for 4 h under standard culture incubation conditions. Following that, 100 μl of 10% SDS in 0.01 M hydrochloric acid solution was added to the wells and the plate returned to the incubator overnight. Optical density reflecting concentration of the formazan dye was determined at 590 nm using a VersaMax plate reader (Molecular Devices, CA, USA).

2.8. Transient transfection of genes using siRNA

A cocktail of three StealthTM siRNA duplexes each for BCRP, aSMase, ceramidase and SPHK1, designed and synthesised by Invitrogen (San Diego, USA), were used for transient inhibition of gene expression in trophoblasts and BeWo cells (Table 2.4).

Gene siRNA sequences aSMase 5’-CCUACCGCCUGGCACAACCUGGUAU-3’

5’-GCCCUGCCGUCUGGCUACUCUUUGU-3’

5’-GAGCUGGAAUUAUUACCGAAUUGUA-3’

Methods

Ceramidase 5’- AACAGCGGCAAUACCCUUCAUUUCC-3’

5’- UUCAGUGUAAGACUGAACAGUCCUG-3’

5’-GGAGUAUUUCUUGGGUGGAACAUAA-3’

SPHK1 5’- CCUACUUGGUAUAUGUGCCCGUGGU-3’

5’-GAGGCUGAAAUCUCCUUCACGCUGA-3’

5’- GCGCUGGCAGCUUCCUUGAACCAUU-3’

BCRP 5’- UAAUGAUGUCCAAGAAGAAGUCUGC-3’,

5’–AUAACCAGCUGAUUCAAAGUAUCCC-3’,

5’-UAAGAUGACACUCUGUAGUAUCCGC-3’

Table 2.4. StealthTM siRNA duplexes used for transient gene silencing

Primary trophoblasts (1.5 million cells/well) were transfected using the ProFection® Mammalian Transfection System (Promega, USA) as outlined below, according to the reverse transfection protocol provided by the manufacturer. Three hours prior to transfection, media was removed from cells and replaced with fresh culture media. For each transfection siRNA and 2X HBS solutions were prepared in separate sterile tubes (Table 2.5). To silence BCRP, all three BCRP siRNAs were combined in equal proportions at a final concentration of 20 nM (optimal concentration calculated from previous work done in BeWo cells using the same siRNA duplexes). Appropriate volumes of siRNA and water were added to Tube 1, followed by addition of CaCl2 and mixed well. Specified amount of 2x HBS was added to Tube 2 and gently vortexed. Contents of Tube 1 were then added drop wise to Tube 2 while vortexing continuously. The combined solution was incubated at room temperature for 30 min prior to being added to cells.

Methods

The CaCl2-siRNA precipitate was added drop wise to each well (150 µl/well for 24- well plates containing 500 µl/well culture media; 24 µl/well for 96-well plates containing 100 µl/well culture media) and plates were swirled to distribute the precipitate evenly over the cells (avoiding localized acidification of cells). Plates were returned to a 37˚C CO2 incubator for 12 h then the transfection process was halted by change of media. Cells were also transfected with an equal concentration of Stealth scrambled siRNA duplexes with a similar GC content to be used as mock-siRNA controls. Functional experiments were carried out 72 h post-transfection as it displayed the most efficient knockdown.

Tube 1 Nuclease free water 66 µl

siRNA 20 nM

CaCl2 9 µl

Tube 2 2X HBS 75 µl

Total volume per well 150 µl

Table 2.5. Volume added per well in a 24 well plate

2.9. Lipid analysis by mass spectrometry

2.9.1. Ceramide and cholesterol extraction

Ceramide and cholesterol analyses were performed on a Thermo Finnigan TSQ Quantum Ultra AM mass spectrometer operating in a multiple reaction monitoring, negative (ceramide) and positive (cholesterol) ionization modes. They were solvent- extracted and measured according to a previously described protocol (Bielawski et al. 2006) as follows: The culture media was removed and cells (2 x 106 cells/well in 6

Methods well plate) were washed with cold PBS x2, before being scraped and collected in 5 ml glass tubes. The cell lysates were centrifuged at 4˚C at 1000 rpm for 5 min, supernatant discarded and the pellet was stored at -80˚C until extraction. A standard curve was prepared from the stock solutions in Chloroform: Methanol (75:25 v/v) by making serial dilutions ranging from 2 µM to 0.075 µM. 50 µl of each standard were added to a fresh 5 ml glass tube each. This was followed by addition of 2 ml PBS to each tube of standard and sample (unknown) after which of internal standard was added per tube and vortexed. Following addition of 2 ml of extraction solvent, the tubes were vortexed, sonicated 3x for 30 sec each and centrifuged for 10 min at 4000 rpm at 4˚C. The upper layer was carefully removed and collected in a new tube, and the extraction was repeated on the bottom layer. The two sets of upper layers were pooled (total volume 4 ml) and evaporated in glass tubes using HetoVac vacuum centrifuge (Heto, Denmark). Reconstitute dried extracts in 200 µl of Chloroform: Methanol (75:25 v/v). Each sample was injected into a Surveyor Autosampler & Surveyor MS pump/TSQ Quantum Ultra AM liquid chromatography/mass spectrometry system, and eluted using isocratic reverse-phase chromatography performed on a Phenomenex Luna 3 µm C18 column (50 x 3 mm) with a 100% methanol mobile phase at a flow rate 800 μl/min. Peaks corresponding to the expected elution profiles of the target analytes and internal standards were identified, collected, analysed and processed using the Thermo Scientific Xcalibur software system (Waltham, USA). Quantitative analysis was based on the calibration curves generated by spiking an artificial matrix with known amounts of synthetic standards and an equal amount of the internal standard. The calibrator/internal standard peak area ratios were plotted against analyte mass to generate the calibration curve. The sample/internal standard peak area ratios were similarly normalized to their respective internal standards and compared with the calibration curves by linear regression. Ceramide concentrations were normalised to total cholesterol levels in the same sample (measured by LC-MS/MS) to control for variability in cell number and extraction losses.

Methods

2.9.2. Sphingosine and S1P extraction

Intracellular Sph and S1P (intracellular and secreted) analysis were performed on a Thermo Finnigan TSQ Quantum Ultra AM mass spectrometer operating in a multiple reaction monitoring, positive ionization mode. SPLs were extracted according to the previously described protocol (Bielawski et al. 2006). The dried extract was reconstituted in 100 l of 1 mM ammonium acetate 0.2% formic acid in methanol and 10 μl injected into a Surveyor Autosampler & Surveyor MS pump/TSQ Quantum Ultra AM liquid chromatography/mass spectrometry system. Isocratic reverse-phase chromatography was performed on a Phenomenex Luna 3 μm C18 column (100 x 2 mm) with a 5 mM ammonium acetate 0.2% formic acid in water/5 mM ammonium acetate 0.2% formic acid in methanol mobile phase at a flow rate 300 μl/min. Peaks corresponding to the expected elution profiles of the target analytes and internal standards were identified, collected, and processed using the Thermo Scientific Xcalibur software system (Waltham, MA, USA). Quantitative analysis was based on the calibration curves generated by spiking an artificial matrix with known amounts of synthetic standards and an equal amount of the internal standard. The calibrator/internal standard peak area ratios were plotted against analyte mass to generate the calibration curve. The sample/internal standard peak area ratios were similarly normalized to their respective internal standards and compared with the calibration curves by linear regression. Sph concentrations were normalised to total cholesterol levels in the same sample (measured by LC-MS/MS) to control for variability in cell number and extraction loses.

Methods

2.9.3. Standard curves and LC-MS/MS parameters for compounds analyzed using LC-MS/MS.

Ceramide and cholesterol Sph and S1P

Mobile Phase 100% methanol (isocratic) 5 mM ammonium acetate 1% formic acid in MilliQ

water/5 mM ammonium acetate 1% formic acid in methanol

Column Luna C18 50x3 mm 3 µm Luna C18 100x2 mm 3 µm (Phenomenex, CA, USA) (Phenomenex, CA, USA)

Column Temperature 30˚C 40˚C

Flow rate 800 µl/min 300 µl/min

Retention times 1.1 min (ceramide) and 1.3 3.8 min (Sph C17), 4.9 min (cholesterol) min (Sph), 5.4 min (S1P C17) and 6.7 min (S1P)

Injection volume 5 µl 10 µl

Wash needle 100% methanol 1% formic acid in methanol

Table 2.6. Liquid chromotography conditions

Methods

Figure 2.1. Standard curves for cholesterol, ceramide, Sph and S1P generated using the Xcalibur software system

Methods

Ceramides Cholesterol

Ion Source Type APCI APCI

Mode Negative Positive

Discharge Current 14 40

Vaporizer Temperature 450 450

Sheath Gas Pressure 50 40

Ion Sweep Gas Pressure 1 0

Auxiliary Gas pressure 3 5

Capillary Temperature 320 250

Sph and S1P

Ion Source Type ESI

Mode Positive

Spray voltage 6000

Methods

Sheath Gas Pressure 60

Ion Sweep Gas Pressure 0

Capillary Temperature 320

Capillary Offset 35

Table 2.7. LC-MS/MS parameters for ceramide and cholesterol (A) and Sph and S1P (B)

Molecule Parent ion Fragment ion Collision Collision Mass Mass Argon Energy Pressure

C16 Ceramide 536.5 280.1 1.5 mTorr 34V

C16 Ceramide- 567.7 310.4 1.5 mTorr 34V D31

Cholesterol 369.3 147.0 1.2mTorr 27V

D6 Cholesterol 375.3 152.0 1.2 mTorr 27V

C17 Sph 286.2 250.2 1.0 mTorr 20V

C18 Sph 300.2 264.2 1.0 mTorr 20V

C17 S1P 366.2 250.2 1.0 mTorr 16V

Methods

C18 S1P 382.3 284.3 1.0 mTorr 16V

Table 2.8. LC-MS/MS instrument methods for ceramide, Sph, S1P and their respective internal controls

2.10. Statistics

For analysis of grouped data from multiple experiments with variable baselines (e.g. immunoblot densitometry), data were first normalised by expressing each data point as a percentage of the sum of all data points from each experiment. The results of multiple experiments were then pooled and analysed by either one-way or two-way ANOVA with Bonferroni test post-hoc using GraphPad Prism Version 5 for Windows (GraphPad Software, San Diego, USA, www.graphpad.com). For two-group comparisons two-tailed paired t-test was used. P<0.05 was considered significant. All data are represented as mean values ± standard error of the mean (SEM).

Results: Chapter 3

CHAPTER 3. TROPHOBLAST DIFFERENTIATION IN VITRO MODELS

3.1. Introduction

Placental cell lines or primary trophoblasts can be used to study the process of villous trophoblast syncytialization (morphological/structural trophoblast differentiation) by monitoring the changes in levels of key genes, enzymes and hormones as well as observing the changes in cellular morphology. Arguably one the most widely cited and investigated biochemical differentiation marker to date is hCG, which is expressed and released by syncytiotrophoblast and in turn contributes towards enhanced syncytial formation and self-synthesis via its autocrine and paracrine actions (Shi et al. 1993; Cronier et al. 1995). However, more recent studies indicated that syncytialization is not a prerequisite for biochemical differentiation i.e. hCG secretion, in line with the theory of the two processes being dissociated (Kao et al. 1988; Orendi et al. 2010). Classical differentiation pathways involve hCG binding to its receptor (hCG/LH receptor) activating adenylyl cyclase, phospholipase C and ion channels, which in turn control levels of cellular cAMP, inositol phosphates, Ca2+ and other second messengers (Gudermann et al. 1992; Hipkin et al. 1992).

The importance of cAMP-mediated signaling in biochemical and morphological trophoblast differentiation has been long appreciated (Strauss III, 1992), and several studies employing cAMP analogs and inhibitors have been able to modulate fusion (Kreyer et al., 1998) and/or biochemical differentiation (Nulsen JC et al 1988). Expression and activation of protein kinase A (PKA) by cAMP has been shown to be central to syncytial formation (Keryer et al. 1998; Knerr et al. 2005). Stimulation of the cAMP-PKA pathway leads to activation of the downstream transcription factor GCM1 (as mentioned in chapter 1). GCM1 is a placenta-specific transcription factor, and a key player in regulating an important balance between trophoblast proliferation and differentiation, a crucial step for normal placental development (Baczyk et al. 2009). Other studied markers of biochemical trophoblast differentiation include proteases, in particular early caspase 8 and executer caspase 3/7, and PLAP. Interestingly, some studies have shown regulation of trophoblast differentiation by

Results: Chapter 3 cAMP-PKA pathway occurring independent to hCG-induced stimulation (Sawai and Azuma 1996; Cronier et al. 1997) (Yang et al. 2003; Lambot et al. 2005).

Based on data from BeWo choriocarcinoma cells, Orendi et al. recently proposed that PKA activation by cAMP was the central driver of fusion, but not hCG secretion (Orendi et al. 2010). Hence the “in vitro trophoblast differentiation model” has recently been revised to discriminate between biochemical differentiation and morphological fusion. Rote et al. proposed that differentiation signals may trigger pathways leading to hCG secretion and thereby regulating the biochemical aspects of differentiation, without directly affecting intercellular fusion (Rote et al. 2010).

This chapter presents the results of studies verifying the reliability of the in vitro trophoblast differentiation model used in this project by assessing levels of above listed differentiation markers combined with morphological assessment of cell fusion. Furthermore, in light of the recent findings by Orendi et al which suggested PKA- regulated syncytialization is independent to hCG secretion (in BeWo cells), the role of PKA in term villous trophoblast differentiation was examined using exogenous small molecule PKA modulators (H89 and FSK).

3.2. Results

3.2.1. Biochemical differentiation markers in primary trophoblasts and BeWo cells

Purity of primary trophoblast cultures was analyzed by immunostaining for presence of cytokeratin-7 (a trophoblast-specific marker) (Figure 3.1A-B) and absence of vimentin (a fibroblast-specific marker) (Figure 3.1C-D). Primary trophoblasts were allowed to differentiate/syncytialize spontaneously over 7 days in culture; cell differentiation was triggered in BeWo cells by FSK treatment. Morphological differentiation (syncytialization) was also assessed by cytokeratin-7 immunostaining on cells fixed at day 1 post isolation (Figure 3.1A) and day 5 of culture (Figure 3.1B). To verify the validity of trophoblast differentiation model, hCG production, GCM1 expression and PLAP activity were assessed in a timely manner (Figure 3.2).

Results: Chapter 3

In primary trophoblasts isolated from term placenta, secretion of hCG increased dramatically on day 2-3 of culture (>10-fold) and then declined rapidly thereafter. In contrast, intracellular hCG levels increased by 20-40-fold from day 2 onwards and remained elevated during the remainder of culture. The amount of secreted and intracellular hCG detected on day 1 was 2.8 µIU/µg and 2.7 µIU/µg, respectively. Secreted hCG from the BeWo (day 1 = 1.56 mIU/µg) cultures showed a similar trend with intracellular hCG levels upregulated by day 2 (>2-fold) and remaining raised as cells differentiated and fused (Figure 3.2A). On the other hand, mRNA expression of the CTB-specific transcription factor GCM1 declined by >50% as trophoblasts differentiated (Figure 3.2B), whereas in BeWo cells no significant decrease was observed during FSK-induced differention and fusion. PLAP activity in trophoblasts showed an upregulation as cells differentiated and fused (Figure 3.2C).

Results: Chapter 3

Figure 3.1. Immunohistochemical identification of primary trophoblasts isolated from the human term placenta stained with the epithelio-specific marker cytokeratin-7 and mesnchymal marker vimentin. (A) CTB cells 24 hours after isolation. (B) Islands of syncytiotrophoblast formed on day 5 after CTB fusion. (C-D) Trophoblasts showed negative staining for vimentin to assess culture purity on days 1 and 5 of culture, respectively. After incubation with the primary antibody, cells were incubated with secondary HRP conjugated-antibody and staining developed using diaminobenzidine (DAB). Nuclei were stained with haematoxylin. Magnification x 40 (A-B), x 20 (C-D); scale bar- 20 µm

Results: Chapter 3

Figure 3.2. Differentiation of primary trophoblasts and BeWo cells in vitro. (A) hCG concentration was measured by ELISA in conditioned media or cellular lysates from primary term trophoblasts or FSK stimulated BeWo cells through 7 days in culture. (B) Expression of the CTB transcription factor GCM1 as determined by quantitative real-time PCR in primary trophoblast and BeWo cultures. (C) PLAP activity in trophoblasts cultured for 7 days. Relative levels are shown as mean ± SEM from n=3-5 experiments. *P≤ 0.05, ***P≤ 0.001 compared to day 1 levels by either one-way or two-way ANOVA with Bonferroni test post-hoc.

Results: Chapter 3

3.2.2. Caspase activity during syncytialization in term villous trophoblasts

Based on the known - albeit controversial - role of caspase 8 in trophoblast differentiation, I also determined levels of caspase 8 activity during differentiation and cell fusion. Caspase 8 activity was initially elevated and then declined from day 4 onwards, albeit none of the changes were significant (Figure 3.3A). Caspase 3/7 activity levels showed a similar trend as trophoblasts differentiated, with an upregulation in activity on day 2, followed by a gradual reduction as trophoblasts syncytialized compared to day 1 (Figure 3.3.B). Activation of caspase 3/7 triggers cleavage of cytokeratin 18 neo-epitope, which can be detected with the M30 antibody to study execution stages of apoptosis. Immunostaining using M30 antibody showed a marked increase in DAB reaction staining for M30-positive cells as trophoblasts progressed from their mononuclear phenotype to their multinucleate syncytialized form (Figure 3.3C).

Results: Chapter 3

Figure 3.3. Caspase 8 and 3/7 activity during trophoblast differentiation in vitro. Activity of caspase 8 (A) and caspase 3/7 (B) was measured for up to 7 days in culture. Caspase 3/7 activity was also assessed by M30 immunostaining on days 1 and 7 of culture (C). Data shown represent mean ± SEM in n=3 experiments, compared to day 1 levels by one-way ANOVA with Bonferroni test post-hoc; scale bar- 20 µm.

Results: Chapter 3

3.2.3. Modulating the PKA pathway during trophoblast differentiation

Spontaneously differentiating primary trophoblasts treated with PKA inhibitor H89 (10 µM) for 3-5 days of culture (Figure 3.4) exhibited significant reduction of hCG secretion (~50%), PLAP activity (~50%) and amount of fusion (~20%), which was calculated from the ratio of multinuclear cells:total cells in culture. Incubation of trophoblasts with 7.5 µM FSK for the same duration resulted in a remarkable ~1,500- fold upregulation in hCG secretion; however, it downregulated PLAP activity by 35% without significantly altering the amount of cell fusion. Treatment dosages used were not toxic to cells.

Results: Chapter 3

Figure 3.4. Effects of H89 and FSK on term trophoblast differentiation. Trophoblasts were treated with 10 µM H89, a PKA inhibitor, or 7.5 µM forskolin (FSK), for 3-5 days during culture. Cell culture media and cell lysates were then collected to assess hCG secretion (day 3) and PLAP activity (day 5), respectively. E- cadherin immunostaining patterns on day 5 were visualized after treatment with H89 and FSK. The ratio of multinuclear cells:total cells in culture was assessed to measure extent of effect on syncytial formation as represented by graphs. Data shown represents mean SEM of n=3 experiments. *P ≤ 0.05, **P≤ 0.01, ***P≤ 0.001 compared to controls by two-way ANOVA with Bonferroni test post-hoc.

Results: Chapter 3

3.3. Discussion

Consistent with the original observations of Kliman et al., the current study showed that in vitro mononuclear cytotrophoblasts aggregate and fuse to form multinucleated structures suggestive of in vivo syncytial formation (Kliman et al. 1986; Douglas and King 1990). In addition, synthesis and secretion of trophoblast-specific hormone hCG was shown to increase during in vitro culture, indicating biochemical differentiation (Knofler et al. 1999; Leitner et al. 2001). A decrease in levels of GCM1 mRNA and an upregulation PLAP activity also confirmed validity of the trophoblast differentiation model in accordance with previous studies (Leitner et al. 2001; Wich et al. 2009). Adhesion of trophoblasts to tissue culture surfaces followed by thorough washing to remove weakly adhered cells and debri ensured the presence of viable, syncytial fragment-free cultures, as well as a pure trophoblast population with negative vimentin staining (Guilbert et al. 2002).

In this study, the decline in hCG secretion levels after day 3 correlates with the downregulation of GCM1 expression in culture, which may indicate a time-dependent regulatory limitation of the fusion process as previously reported (Alberts et al. 1993; Khan et al. 2000; Wich et al. 2009). Differences in hCG secretion trends between trophoblasts and BeWo cells during cell differentiation may reflect potentially different mechanisms involved in this process. This might help explain the differential effects of various modulators on these cells types, as discussed in subsequent chapters in this thesis. Therefore, caution should be exercised when interpreting differentiation- related data in BeWo cells using hCG secretion as a marker. For the first time, to the best of our knowledge, the present data highlights a critical difference in trends between secreted hCG versus intracellular hCG levels during spontaneous trophoblast differentiation. It is essential to take this into account when investigating hCG-related mechanisms in trophoblast differentiation studies as it may indicate intrinsic differences in function between the two hCG pools. In addition, it has been shown the syncytialization is not a prerequisite for biochemical differentiation i.e. hCG secretion, and the current study is in agreement with those findings (Kao et al. 1988; Orendi et al. 2010). In fact, it has been suggested that biochemical and morphological differentiation are parallel independent events, and this thesis will provide further evidence towards this notion.

Results: Chapter 3

Until recently, it was believed that during the initial stages of CTB differentiation and fusion into syncytiotrophoblast, early and reversible stages of the apoptotic cascade were involved. These include the cleavage of caspase 8 substrate alpha-fodrin accompanied by the translocation of PS from the inner to outer side of the plasma membrane (Huppertz et al. 1998; Huppertz et al. 1999). It was commonly accepted that these pre-apoptotic stages were prerequisites for syncytial fusion, since blocking caspase-8 activity (detectable in CTB cells prior to fusion) prevented PS externalization and subsequent syncytialization (Adler et al. 1995; Vogt et al. 1997; Huppertz et al. 1999; Black et al. 2004; Huppertz and Kingdom 2004; Leslie et al. 2005). However, these existing differentiation-apoptosis dogmas have been challenged by recent studies postulating the proform and proteolytic active form of caspase 8 may potentially have different roles, with the former contributing to early biochemical differentiation and the latter cleaving fodrin at the last steps of fusion (Guilbert et al. 2010; Rote et al. 2010). Other studies have published data more recently indicating the absence of a relationship between caspase activation in general (and caspase 8 in particular), and biochemical trophoblast differentiation and syncytial formation (Guilbert et al. 2010).

The current study to some extent strengthens the findings of Guilbert et al. by showing the absence of significant changes in activity of early and downstream effector caspases 8 and 3/7 for up to 7 days in culture during biochemical trophoblast differentiation and syncytialization (Guilbert et al. 2010). Guilbert and colleagues used caspase inhibitors in primary trophoblasts from term placentae to inhibit caspase activation, and demonstrated that that inhibition had no effect on trophoblast differentiation, thereby ruling out the role of activated caspases in the trophoblast differentiation process (Guilbert et al. 2010). Interestingly, caspase 3/7 data from the current study showed a similar trend to findings by Yusuf et al., that reported caspase 3 activity declined gradually post-isolation (Yusuf et al. 2002). Their data also suggested that activation of these caspases occurs while CTBs are in their mononuclear state, with subsequent intercellular differentiation and fusion progression in the absence of further caspase activation (Yusuf et al. 2002). The lack of a unifying pattern defining the non-apoptotic role of caspases in differentiation and cell division may be due to the specific role of caspases varying considerably among different cell types (Maelfait and Beyaert 2008). Furthermore, unlike cells undergoing apoptosis, differentiating trophoblasts lose membrane asymmetry only transiently during cell

Results: Chapter 3 fusion without terminal progression of the apoptotic cascade (Evseenko et al. 2007c). Mechanisms underlying this process remain poorly understood.

Like primary CTBs, BeWo cells transform to a differentiated (hCG secreting) phenotype when treated with FSK, which stimulates cAMP production and PKA activation leading to cell fusion (Wice et al. 1990; Knerr et al. 2005; Rote et al. 2010). Interestingly, as this work was being carried out, a study was published in BeWo cells suggesting that cAMP-dependent PKA activation was primarily involved in biochemical differentiation (hCG secretion), whereas syncytialization was regulated by activation of a non-PKA-dependent pathway (Yoshie, Kaneyama et al. 2010). On the other hand, another recent study in BeWo cells, reported regulation of morphological differentiation via the cAMP-PKA pathway independent of upregulation of hCG, concluding that hCG was not a reliable differentiation marker (Orendi et al. 2010; Yoshie et al. 2010). The present data in term trophoblasts does not support these hypotheses, as explained above. In accordance with previously cited literature, the current study showed a significant downregulation in biochemical (hCG secretion) and morphological differentiation in response to PKA inhibition (Keryer et al. 1998; Lin et al. 2000).

It should be recognized that, at the concentration used in these studies (10 μM), H89 has been shown to inhibit several kinases with greater potency than PKA, including ERK1/2 and Akt (Murray et al, 2008). ERK1/2 is an established regulator of trophoblast differentiation, and as shown in Chapter 5 of this thesis, Akt is also associated with trophoblast differentiation. H89 also alters intracellular Ca2+ levels which may regulate hCG secretion directly or indirectly, independent of PKA (Hussain et al. 1999; Davies et al. 2000; Palacois et al. 2007). However, in trophoblast differentiation studies, H89 has been used to investigate PKA-related regulation of trophoblast differentiation, and our data are in agreement with these findings (Keryer et al. 1998; Lin et al. 2000). It is difficult to explain the discrepancies between cAMP- PKA regulated cell differentiation and fusion in BeWo cells and term trophoblasts, but our studies suggest that the cAMP-PKA pathway is involved in both the biochemical and morphological aspects of trophoblast differentiation and fusion.

It is interesting to note that although FSK upregulated hCG secretion and cell fusion in trophoblasts, it had a negative effect on PLAP activity. Lack of reliability of PLAP as

Results: Chapter 3 a fusion marker has been raised previously due to its expression in mononuclear CTBs; however, my data from spontaneously differentiating trophoblasts show a significant upregulation in PLAP activity as expected (Leitner et al. 2001). Therefore, it is plausible that the downstream targets of FSK-activated cAMP in trophoblasts vary to those of BeWo cells, thereby resulting in differential effects on PLAP activity.

To summarize these findings, this study highlights the possibility of differential functions of different pools of hCG (secreted vs. intracellular) based on their varied trends as trophoblasts differentiated, aggregated and fused. Differences in hCG secretion patterns between primary trophoblasts and BeWo cells is an important reminder to assess data in choriocarcinoma cells with caution when simulating primary cell culture models. Although choriocarcinoma-derived BeWo cells have been extensively used in vitro to study villous trophoblast differentiation, it should be appreciated that these cells are transformed and their differentiation, syncytial formation and protein expression patterns differ from primary trophoblasts (Filla and Kaul 1997; King et al. 2000). The absence of a correlation between caspase activity and syncytialization is in line with recent studies argueing against the involvement of these proteases in the process. The current data agrees with conventional studies identifying the cAMP-PKA pathway as a regulator of functional and morphological differentiation. The divergent effects of FSK on hCG production and PLAP activity in this study may support the notion that at least some aspects of biochemical and morphological trophoblast differentiation can be dissociated (Kao et al. 1988; Johnstone et al. 2005; Al-Nasiry et al. 2006; Rote et al. 2010).

Results: Chapter 4

CHAPTER 4. BCRP TRANSPORTER, SPLs AND TROPHOBLAST DIFFERENTIATION

4.1. Introduction

BCRP/ABCG2, also referred to as “placenta-specific ABC transporter” or “ABC transporter in placenta” (ABCP), is most highly expressed in the placenta at both mRNA and protein levels (Mao 2008). This transporter is highly expressed on the apical membrane of the syncytium, and as such is one of the major placental drug efflux transporter contributing towards MDR, along with MDR1/ABCB1 (Evseenko et al. 2006). Although, there is contradicting literature on the expression patterns of BCRP during gestation, the general consensus remains that BCRP plays a crucial protective role by expelling metabolites, drugs and xenobiotcs especially during mid- gestational ages (Mathias et al. 2005; Meyer zu Schwabedissen et al. 2006; Yeboah et al. 2006) (Yasuda et al. 2005; Kalabis et al. 2007; Mao 2008). Considering the variation in expression patterns of the other key drug efflux transporters, MDR1 and MRP1, in the syncytial apical membrane over gestation, it is likely that collectively these transporters provide a protective mechanism for the fetus throughout pregnancy by their differential expression (Mao 2008).

Transport of substrates by BCRP into the fetal circulation has also been suggested by its detection in the endothelial cells lining the fetal capillaries (Ceckova et al. 2006; Evseenko et al. 2006). BCRP transports a wide range of substrates, including organic anions, endogenous lipid substrates (e.g. steroids and their conjugates), chemotherapeutic agents, anti-hypertensives, antibiotics and anti-diabetic drugs (Woehlecke et al. 2003; Choudhuri and Klaassen 2006; Mao 2008). In addition, this placenta-specific ABC transporter also provides protection from other endogenous and exogenous stress factors including proinflammatory mediators, and has been associated with the regulation of intracellular toxic metabolites, such as prostaglandins, SPLs and porphyrins in normal and cancer cells (Johnstone et al. 1999; Pallis and Russell 2000; Paumi et al. 2003; Aye et al. 2009).

Due to high expression levels in the placenta, speculations of an alternative role of BCRP besides that of a drug effluxer have surfaced. It is associated with regulation of trophoblast survival during pregnancy complications as it confers resistance to

Results: Chapter 4 hypoxia by pumping out toxic products from the cytoplasm (Krishnamurthy et al. 2004). During biochemical and morphological trophoblast differentiation, an upregulation in BCRP expression in vitro has also been demonstrated (Evseenko et al. 2006). Silencing BCRP expression in BeWo cells undergoing FSK-induced differentiation caused diminished cell differentiation and an increase in apoptosis (Evseenko et al. 2007c). Moreover, silencing this protein enhanced accumulation of endogenous ceramides in cells undergoing cytokine-induced stress and reduced the ability to retain PS in the inner leaflet of the plasma membrane in non-stressed trophoblasts (Evseenko et al. 2006; Evseenko et al. 2007b). These studies imply an involvement of BCRP in the regeneration of the placental syncytial epithelium. BCRP may have an as-yet undefined role as a differentiation/survival factor in the trophoblast differentiation process, as it displays protective functions in trophoblasts against apoptosis induced by TNF-α and short chain ceramides (Evseenko et al. 2007a). However, the exact underlying mechanisms await elucidation (Evseenko et al. 2007a; Evseenko et al. 2007c).

In the current chapter, we have used term trophoblasts to partly replicate the work of Evseenko et al. in BeWo cells to investigate whether BCRP plays a similar role in trophoblast differentiation regulation as it does in FSK-stimulated differentiating BeWo cell lines (Evseenko et al. 2007c). We further explored whether key membrane component SPLs are involved with the pro-differentiation function of BCRP. This study was performed to help further expand upon our knowledge of the interaction and importance of cellular lipids and ABC transporters with respect to the formation of the syncytiotrophoblast layer, essential for placental development and a normal pregnancy. This may help to identify potential causes of placental deficits through dysregulation of SPL synthesis or metabolism and/or interference with ABC function or expression in the syncytium.

Results: Chapter 4

4.2. Results

4.2.1. Effect of expression knockdown and functional inhibition of BCRP on biochemical trophoblast differentiation

Expression patterns of BCRP transporter were investigated over 7 days of culture in trophoblasts isolated from term placenta. An upregulation in BCRP expression was observed on day 2 by ~2-fold compared to day 1, with a decline thereafter by 30%; levels then remained constant as the cells achieved terminal differentiation (Figure 4.1A). Trophoblasts were reverse transfected with a cocktail of three StealthTM BCRP siRNA duplexes (final concentration 20 nM each) and BCRP protein expression was determined by immunoblotting. Transfection conditions were based on successful experiments carried out in trophoblast-like BeWo cells as previously reported (Evseenko et al. 2007c). A significant decrease in BCRP protein was detected at 48 and 72 h after transfection by approximately 59% and 71%, respectively (Figure 4.1B). In control experiments, cells that transfected with the same concentrations of the StealthTM scrambled siRNA with the same GC content exhibited no changes in BCRP expression.

Suppressing BCRP expression had no significant effect on hCG secretion (Figure 4.2A) and caspase 8 activity (Figure 4.2C) after 72 h post-transfection. Similarly, inhibiting BCRP activity using 2.5 µM FC had no effect on hCG accumulation and caspase 8 activity (Figure 4.2 B, D respectively). To assess the level of BCRP function in BCRP-silenced and -inhibited trophoblasts, cell viability was measured in response to exposure to MTX, a high affinity cytotoxic BCRP substrate, for 72 h as per previous studies(Evseenko et al. 2007a). MTX had a significantly toxic effect, causing a 50% decrease in cell viability (Figure 4.3A, B). While inhibition of BCRP activity enhanced cytotoxic effects of MTX by approximately 50% (Figure 4.3B), downregulation of BCRP expression did not significantly enhance MTX cytotoxicity (Figure 4.3A).

Results: Chapter 4

Figure 4.1. Suppression of BCRP expression in term trophoblasts using transient transfection with StealthTM BCRP siRNA. (A) Changes in BCRP protein expression in culture, coinciding with alterations in biochemical and morphological differentiation (previous chapter), were detected using Western blotting. (B) Control cells were transfected with StealthTM BCRP (20 nM final concentration) or scrambled siRNA at the same final concentration; immunoblots showing silencing of BCRP expression. Data are presented as mean ± SEM; n=5-7. **P<0.01 vs day 1 (A); ***P<0.001 compared to control cells (B) by either one-way or two-way ANOVA with Bonferroni test post-hoc.

Results: Chapter 4

Figure 4.2. Effect of BCRP silencing and inhibition on trophoblasts during spontaneous differentiation. (A, B) hCG secretion levels in response to BCRP siRNA and functional inhibition by FC after 72 h, respectively. (C, D) Caspase 8 activity was assessed post-siRNA-mediated transfection and functional inhibition by FC after 72 h, respectively. Data are presented as mean ± SEM compared by two- tailed paired t-test; n= 5.

Results: Chapter 4

Figure 4.3. Assessment of BCRP function and trophoblast viability. (A) Cell viability in response to BCRP silencing and a measure of the cytotoxic effects of MTX, a high affinity BCRP substrate. (B) Effect of BCRP inhibition, with FC, on cell viability and assessing loss of function using MTX. Data are presented as mean ± SEM; n= 4. **P<0.01, ***P<0.001 compared to controls by by one-way ANOVA with Bonferroni test post-hoc.

Results: Chapter 4

4.2.2. Regulation of SPL enzymes and metabolites in response to BCRP silencing

The downstream effects of BCRP silencing on SPL levels and expression of SPL- modifying enzymes were measured for over 72 h post-transfection of cultured term trophoblasts during spontaneous differentiation and syncytialization. Protein expression of key SPL metabolic enzymes, aSMase (Figure 4.4A), ceramidase (Figure 4.4B), CERK (Figure 4.4C), SPHK1 (Figure 4.4D) and S1PP2 (Figure 4.4E), remained significantly unaltered in BCRP-silenced trophoblasts. Minor changes in aSMase and ceramidase expression were observed in treated cells compared to controls; however, these changes were not statistically significant.

Levels of SPL enzyme metabolites - ceramide and Sph - were also assessed using LC- MS/MS in lipid extracts of post-transfected trophoblasts. Silencing of BCRP resulted in significant ceramide accumulation 24 h post-BCRP transfection during the differentiation and fusion process (Figure 4.5A; P<0.05). On the other hand, Sph levels remained unchanged in response to BCRP silencing (Figure 4.5B).

Effects on BCRP expression were also investigated in response to exogenous SPL administration C8 ceramide (Figure 4.6A), Sph (Figure 4.6B) and S1P (Figure 4.6C) in order to determine whether these bioactive lipids regulate BCRP levels. No significant changes were observed in protein expression of this transporter in response to these bioactive lipids during trophoblast differentiation in vitro. Concentrations of exogenous SPLs employed were as cited previously (Johnstone et al. 2005; Evseenko et al. 2007a; Morales et al. 2007; Yoon et al. 2009).

Results: Chapter 4

Figure 4.4. Expression of key SPL enzymes aSMase (A), ceramidase (B), CERK (C), SPHK1 (D) and S1PP2 (E) during trophoblast differentiation and in response to BCRP silencing. Data are presented as mean ± SEM compared by two- way ANOVA with Bonferroni test post-hoc to control cells in n=4 experiments.

Figure 4.5. Effect of BCRP StealthTM siRNA on endogenous levels of C16 ceramide (A) and Sph (B) measured by LC-MS/MS. SPL levels were normalized to cellular cholesterol. Data are presented as mean ± SEM compared by two-way ANOVA with Bonferroni test post-hoc to control cells in n=4 experiments.

Results: Chapter 4

Figure 4.6. BCRP expression in response to short chain ceramide (A), Sph (B) and S1P (C). Data are presented as mean ± SEM compared by two-way ANOVA with Bonferroni test post-hoc to control cells in n=4 experiments.

Results: Chapter 4

4.3. Discussion

The current study argues against the hypothesis that a relationship exists between BCRP and biochemical trophoblast differentiation regulation in vitro (Evseenko et al. 2007c). Discrepancies with previous findings may be largely due to the different cell type and differentiation models employed to investigate this hypothesis. The choice of BeWo cells in former studies, albeit a popular alternative to trophoblasts due to their ease of transfection, has significant limitations as mentioned in the previous chapter. The elevated expression levels of BCRP as trophoblasts commence differentiation in culture may protect trophoblasts from progression of apoptosis during the period of caspase activation and loss of the plasma membrane asymmetry associated with early phases of cell apoptosis (Evseenko et al. 2007a). Interestingly, although a 70% downregulation of BCRP expression was achieved in the transfection studies of the current study, it failed to augment the cytotoxic effects of MTX in BCRP-silenced cells, unlike BCRP inhibition which was effective. This may be because BCRP is expressed so abundantly in the human placenta that a 70% knockdown in expression is not significant enough to alter its biological functions. Despite the functional downregulation in BCRP’s protective effects observed in trophoblasts treated with FC, this was ineffective in altering levels of hCG secretion. In addition, the absence of an effect of caspase 8 in BCRP inhibited and silenced cells questions its protective functions via the caspase cascade. Thus, the present findings suggest that BCRP does not have a role in biochemical trophoblast differentiation.

ABC proteins actively participate in maintaining lipid asymmetry across cellular membranes, including the plasma membrane (Sietsma et al. 2001). Studies have shown an active role for these proteins in transport and regulation of a variety of endogenous compounds, including SPLs (Aye et al. 2009). A number of recently identified members, particularly from ABCG subfamilies, have been identified as key molecules in the regulation of cellular lipid transport and whole body lipid homeostasis (Aye et al. 2009). Moreover, studies have shown that the function of many of these proteins is, in turn, dependent on the membrane lipid milieu in which they reside. Membrane lipids are involved in regulating the functional activity and/or expression of membrane proteins, in particular ABC transporters, as they are an integral part of the plasma membrane (Van Helvoort et al. 1996; Kok et al. 2000).

Results: Chapter 4

A considerable amount of evidence suggests that components of the SPL biosynthetic/metabolic pathways are regulators of ABC protein functions. In line with these findings, accumulation of endogenous long chain ceramides was observed in BCRP-silenced BeWo cells (Evseenko et al. 2007c). Similarly, in the current study using primary trophoblast cultures modestly elevated ceramide levels were observed, which rapidly returned to basal levels 24 h later. However, this data should be interpreted with caution since the increase in ceramide accumulation occurred prior to significant reduction in BCRP protein expression. In addition, expression of ceramide synthesis/metabolic enzymes remained unaltered in transfection studies. Hence, our current data showed a lack of correlation between SPL levels, expression of their hydrolytic enzymes and BCRP (Mitra et al. 2006; Aye et al. 2007; Aye et al. 2009).

In conclusion, this study did not support a role for BCRP in biochemical/hormonal trophoblast differentiation in the human placenta. Results derived using BeWo cells to determine the role of BCRP during trophoblast differentiation in an earlier study were not reproducible in this thesis chapter using primary cells (Evseenko et al. 2007c). Therefore, as mentioned in the previous chapter, the use of this choriocarcinoma cell line should be used with extreme caution when simulating trophoblast characteristics. It should be appreciated that BeWo cells exhibit different expression and functional patterns to trophoblasts. Furthermore, these current findings reveal a lack of association between SPLs (ceramide and Sph) and BCRP during trophoblast differentiation and syncytialization.

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Results: Chapter 5

CHAPTER 5. ROLE OF CERAMIDE BIOSYNTHESIS AND METABOLISM IN THE REGULATION OF TROPHOBLAST DIFFERENTIATION AND SYNCYTIALIZATION

5.1. Introduction

SPLs have been traditionally considered to be the building blocks of the plasma membrane, but are now known to play a key role in regulating several fundamental cellular biological activities including cell differentiation, survival and proliferation (Morales et al. 2007). An increasing number of studies have described a variety of downstream signaling pathways activated by SPLs that regulate many different biological cell functions, in particular differentiation (Mao and Obeid 2008). One of the most highly studied SPL mediators is ceramide, an N-acylsphingosine produced primarily from SM or Sph and at the centre of SPL biosynthesis and degradation pathways (Hannun and Obeid 2002). Ceramide has been studied in various cell types mainly for its role in cellular differentiation and apoptosis, and has been shown to exert various biological actions depending on its intracellular localization, downstream targets and cell state (Hannun and Obeid 2002; Futerman and Hannun 2004; Sun et al. 2008). It plays a key role in cell signaling, and forms the hydrophobic backbone of complex SPLs (Cremesti et al. 2002). The key pathways involved in the synthesis of ceramide include the de novo synthetic pathway (mediated by CerS), the SMase-mediated pathway (via acid or neutral SMases), and the salvage pathway (catabolism of complex glycosphingolipids into Sph, which is recycled via reacylation to produce ceramide via) (Futerman and Hannun 2004). Members of the ceramidase family metabolize ceramide to produce sphingosine, which can be reconverted back to ceramide via CerS, or phosphorylated by SPHK to generate S1P, a highly potent and biologically active extracellular mediator (Futerman and Hannun 2004). Ceramide can also be phosphorylated by CERK to C1P which has pro-inflammatory actions (Gangoiti et al.).

SPL homeostasis is highly complex and regulated by numerous extracellular stimuli (Okazaki et al. 1989). It can be regulated on a cell-specific basis, or via signals produced by controlling allosteric mechanisms, post-translational modifications or expression of enzymes (Futerman and Hannun 2004). A multiphasic increase in

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Results: Chapter 5 intracellular levels of ceramide and its metabolites is observed in response to agonists such as cytokines and growth factors, which cause multiple changes in different enzymes at different times in different subcellular locations (Bourteele et al. 1998). In addition, studies have demonstrated differential lipid accumulation and subsequent cellular responses as a result of alterations in the concentrations of the same agonist (Nikolova-Karakashian et al. 1997). Net ceramide content depends not only on the relative levels of activities of ceramide metabolizing enzymes and other metabolically related bioactive lipids (for example S1P) which may oppose the actions of ceramide, but on the activity of the major ceramide biosynthetic pathways (Jarvis et al. 1994; Cuvillier et al. 1996; Auge et al. 1999).

Despite extensive information from other cellular systems on regulation of differentiation and apoptosis by ceramide, the production and functions of this ubiquitous SPL in the human placenta remain essentially unexplored. In light of the well established role of ceramide in regulation of cell differentiation and apoptosis, we hypothesized that ceramide is likely to be involved in regulation of the trophoblast differentiation process through changes in either its biosynthesis or metabolism. To explore this hypothesis, we assessed changes in various ceramide-related metabolic enzymes during trophoblast differentiation in vitro in correlation with intracellular ceramide levels. We also manipulated concentrations of exogenous ceramide and altered internal ceramide levels via inhibition of key ceramide metabolic enzymes to study their effects on biochemical and morphological differentiation.

5.2 Results

5.2.1. Expression levels of endogenous aSMase, ceramidase and CERK during spontaneous biochemical and morphological trophoblast differentiation.

We determined protein expression levels of three key enzymes involved in the synthesis and metabolism of ceramide during differentiation of trophoblasts in vitro: aSMase, which liberates ceramide from SM in the plasma membrane, endosomes and Golgi (Futerman and Hannun 2004); CERK which phosphorylates ceramide to produce ceramide-1-phosphate (C1P) (Futerman and Hannun 2004), and acid

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Results: Chapter 5 ceramidase which degrades ceramide to form Sph (Futerman and Hannun 2004). As shown in Figure 5.1, aSMase expression was markedly upregulated during the process of biochemical and morphological differentiation between days 2 and 4 of culture (P=0.01), declining slightly thereafter but remaining elevated as trophoblasts progress to become terminally differentiated (Figure 5.1A). CERK protein expression rose modestly but progressively through culture reaching levels around 40% higher in fused cells on day 5 (Figure 5.1B; P=0.0205). On the other hand, ceramidase expression showed a much more dramatic 5-fold increase on day 7 of culture (Figure 5.1C; P=0.0005). Collectively, these changes would predict early phase increased production of ceramide coinciding with advanced biochemical differentiation in vitro followed by increased ceramide metabolism associated with syncytialization.

SPLs were extracted from term primary trophoblasts on days 1, 3 and 5 of culture and LC-MS/MS employed to determine C16 ceramide levels, this being one of the most abundant ceramide species in mammals (Cremesti et al. 2002). In addition, C16-24 ceramides were previously analysed in BeWo cells and it was found that C16 was by far the dominant form (Evseenko et al. 2007), hence the decision to study C16 ceramide in this study. Intracellular C16 ceramide levels peaked on day 3 of culture, coinciding with elevated hCG secretion levels, then declined to basal levels by day 5 (Figure 5.2). Ceramide concentrations were compared to total cholesterol levels in the same sample. Mean amount of intracellular C16 ceramide on day 1 was 1299.9 mg.

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Figure 5.1. Expression levels of ceramide synthetic and metabolic enzymes during trophoblast differentiation in vitro. Using Western blotting, protein expression patterns of aSMase (A), CERK (B) and ceramidase (C) from term trophoblasts were studied through 7 days in culture during spontaneous biochemical and morphological differentiation. Data shown represent mean ± SEM from n=5 sets of cultures. *P≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001 compared to day 1 levels by one-way ANOVA with Bonferroni test post-hoc.

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Figure 5.2. Endogenous levels of ceramide during trophoblast differentiation in vitro. LC-MS/MS was used to identify endogenous levels of C16 ceramide (normalized to cholesterol) in differentiating trophoblasts. Data shown represent mean ± SEM in n=7 experiments; *P ≤ 0.05 compared to day 1 levels by two-way ANOVA with Bonferroni test post-hoc.

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5.2.2. Regulation of trophoblast differentiation by ceramide and its synthesis/metabolic enzymes

To investigate the role of ceramide in trophoblast differentiation, cells were exposed to cell-permeable short chain C8 ceramide (Morales et al. 2007), exogenous aSMase (Mathias et al. 1993), acid ceramidase inhibitor B13 (Selzner et al. 2001; Raisova et al. 2002; Samsel et al. 2004), neutral ceramidase inhibitor D-erythro-MAPP (Gangoiti et al. 2010) and CerS inhibitor fuminosin B1 (Gangoiti et al. 2010) daily for up to 72 h (Figure 5.3).

Figure 5.3. Illustration of pathways targeted with exogenous ceramide modulators and enzyme inhibitor to investigate their functions in regulating biochemical and structural trophoblast differentiation.

Cell viability in response to these exogenous manipulations was assessed every 24 h and none of the treatments altered cell viability in a significant manner (Figure 5.4). Exogenous administration of 10 µM C8 ceramide resulted in enhanced hCG secretion that achieved significance at 72 h (95% above control; P<0.05), consistent with an

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Results: Chapter 5 upregulation of hCG secretion levels (Figure 5.5A). Cells treated with 0.01 U/ml aSMase, which hydrolyses SM to ceramide (Figure 5.5B), exhibited increased hCG secretion by ~80% at the 48 h time point. Treatment with 25 µM B13 to inhibit ceramide metabolism resulted in increased hCG levels of ~110%, but only at the 72 h time point. This suggests that ceramidase may only be biologically significant at the later time point (Figure 5.5C). Neutral ceramidase inhibitor D-erythro-MAPP (10 µM) and CerS inhibitor fuminosin B1 (10 µM) had no significant effect on secretion levels of hCG during biochemical trophoblast differentiation in vitro (Figure 5.5D,E). Intracellular levels of hCG were also determined after exposure to these compounds. Besides B13, which had a similar effect on hCG expression as it did on secretion, none of the other treatments affected hCG expression significantly, although the trends were similar to hCG secretion studies (Figure 5.6A, B, D, E). B13 caused a ~95% upregulation in hCG expression after 72 h of culture (Figure 5.6C).

To determine whether any of the ceramide modulators and ceramidase inhibitor used had direct effects on hCG secretion that may be independent of their effects on biochemical differentiation, trophoblast cultures were exposed to modulators for 24 h on day 4 of culture (i.e. after hCG secretion levels in controls had declined). As shown in Figure 5.7, no significant changes in hCG secretion were observed with C8 ceramide (Figure 5.7A), aSMase (Figure 5.7B) or B13 (Figure 5.7C) compared to controls. A trend towards a decline in hCG secretion was seen in response to B13 which did not reach significance. Further use of D-erythro-MAPP and fuminosin B1 in differentiation studies was discontinued as they had no effect on hCG production, the predominant differentiation marker in this study.

GCM1, a CTB transcription factor that triggers expression of the fusion protein syncytin-1, exhibited a small (~25%) decline in mRNA expression (Figure 5.8A) in response to C8 ceramide, consistent with advanced structural differentiation of mononuclear cells. Exogenously administered aSMase had no effect on GCM1 expression (Figure 5.8B). Inhibition of ceramidase activity showed a strong trend towards decreased GCM1 expression, but did not reach statistical significance (Figure 5.8C). The activity of other commonly used biochemical differentiation markers such as caspase 8 and PLAP also remained unchanged after exposing trophoblasts for 72 h to C8 ceramide (Figures 5.8D, G), aSMase (Figures 5.8E, H) and B13 (Figures 5.8F, I), respectively.

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Cell fusion was assessed immunocytochemically using E-cadherin staining to visualize cell-cell boundaries and counting the relative numbers of multinuclear cells (2 or more nuclei per cell) vs. total number of nuclei (Figure 5.9B-D) and compared to control cells (Figure 5.9A). Cadherins represent another significant family of membrane proteins associated with trophoblast differentiation and syncytial fusion, and can be broadly divided into four subgroups: the classical (type I) and closely related type II cadherins, desmosomal cadherins and protocadherins (Aplin et al. 2009). Of the classical and non-classical cadherins, the most commonly studied ones in trophoblast differentiation include E-cadherin and cadherin-11, respectively (Aplin et al. 2009). Syncytial fusion was investigated following administration of C8 ceramide, aSMase or B13. No significant changes in cell fusion rates were detected over 5 days in culture in response to C8 ceramide (Figure 5.9E) and aSMase (Figure 5.9F). However, inhibition of ceramidase activity resulted in an inhibition of fusion as reflected by reduced multinuclear:total nuclei ratios (Figure 5.9G) (P<0.05).

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Figure 5.4. Assessment of toxicity of various SPL mediators or enzyme inhibitors in term trophoblasts. Cells were treated with C8 ceramide (A), aSMase (B), ceramidase inhibitors B13 (C) and D-erythro-MAPP (D), and CerS inhibitor fuminosin B1 (E) for 72 hr and then viability was determined by MTT assay. Data shown represent mean ± SEM of n=5 experiments, analysed by two-way ANOVA with Bonferroni test post-hoc.

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Figure 5.5. Regulation of biochemical trophoblast differentiation by ceramide, aSMase and B13. Exogenous administration of 10 µM C8 ceramide (A), 0.01 IU/ml aSMase (B) and 25 µM B13 (C), 10 µM D-erythro-MAPP (D) and 10 µM fuminosin

B1 (E) and was carried out to assess their effects on hCG secretion. Data shown represent mean ± SEM of n=5 experiments. *P ≤ 0.05, **P ≤ 0.01 compared to controls for each treatment at respective time points by two-way ANOVA.

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Figure 5.6. Differential effect of ceramide, aSMase and B13 on hCG expression. Spontaneously differentiating trophoblasts were treated with C8 ceramide (A), aSMase (B), ceramidase inhibitors B13 (C), D-erythro-MAPP (D) and fuminosin B1 (E) for 72 h to assess the effects on hCG protein expression. Data shown represent mean ± SEM of n=5 experiments. *P ≤ 0.05 compared to controls for each treatment at respective time points by two-way ANOVA with Bonferroni test post-hoc.

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Figure 5.7. Lack of post-differentiation effects of ceramide, aSMase and B13. To exclude a direct effect of ceramide modulators and B13 on hCG secretion independent of differentiation, trophoblasts were treated with C8 ceramide (A), aSMase (B) and B13 (C) on day 4 of culture after cells had undergone differentiation; cultured media were collected after 24 h for analysis. Data shown represent mean ± SEM of n=4 experiments.

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Figure 5.8. Ceramide, aSMase and B13 have disparate effects on biochemical differentiation markers. Exogenous administration of C8 ceramide (A, D, G), aSMase (B, E, H) and B13 (C, F, I) was carried out to assess effects on GCM1 expression, caspase 8 and PLAP activity during trophoblast differentiation in culture, respectively. Data shown represent mean ± SEM of n=5 experiments, compared to controls by two-way ANOVA with Bonferroni test post-hoc.

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Figure 5.9. Morphological differentiation in response to ceramide modulators and ceramidase inhibitor. E-cadherin immunostaining patterns were visualized after treatment with C8 ceramide (B), aSMase (C) and B13 (D) for 72 h and compared to control cells (A). The ratio of multinuclear cells:total cells in culture was then assessed to measure extent of effect on syncytial formation as represented by graphs (E-G). Data shown represent mean ± SEM of n=3 experiments; *P ≤ 0.05 compared to controls by t-test; scale bar- 20 µm. Multinuclear cells are indicated by arrows.

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5.2.3. Effects of silencing of aSMase and ceramidase expression

Attempts to silence expression of aSMase or ceramidase expression in primary term trophoblasts using a calcium phosphate-based transfection protocol were not successful despite the fact that I have successfully employed this same approach to silence expression other genes in these cells in Chapter 4. Although a ~75% downregulation in aSMase mRNA expression was observed 24 h post-transfection, the effect was abolished within the next 24 h (Figure 5.10A). Likewise, a significant 50-60% decrease was detected in ceramidase mRNA expression at 24 h and 48 h time points after trophoblasts were transfected with StealthTM siRNA; however, the effect was short lived (Figure 5.10B). Therefore, these studies were performed in the BeWo choriocarcinoma cell model to further explore the effects of modification of endogenous ceramide biosynthesis and metabolism on cell differentiation.

Expression of aSMase in BeWo cells was suppressed by siRNA-mediated transfection by ~40-70% up to 72 h in culture (Figure 5.10C). However, this had no significant effect on hCG secretion (Figure 5.11A) or GCM1 expression (Figure 5.11B). Similarly, down-regulation of ceramidase expression (Figure 5.10D) using siRNA also failed to influence BeWo hCG secretion (Figure 5.11C) and GCM1 expression (Figure 5.11D) during biochemical differentiation. To determine whether differentiation in BeWo cells showed the changes in SPL enzyme expression similar to those seen in primary trophoblast cultures, we determined levels of ceramidase, aSMase and CERK proteins during BeWo differentiation by immunoblotting (Figure 5.12). Levels of these proteins did not change significantly throughout the culture period, failing to recapitulate the differentiation-specific changes observed in primary trophoblast.

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Figure 5.10. Efficiency of siRNA-mediated transfection in primary trophoblasts compared to immortalized BeWo cells. Administration of StealthTM siRNA using calcium phosphate to transfect cells and suppress expression of aSMase and ceramidase in trophoblasts (A and B) and BeWo cells (C and D). Data shown represent mean ± SEM of n=3 experiments. *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001 compared to controls by two-way ANOVA with Bonferroni test post-hoc.

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(%)

hCG secretion hCG hCG secretion hCG

Figure 5.11. Silencing of aSMase and ceramidase expression in BeWo cells using StealthTM siRNA. Levels of hCG secretion (A,B) and GCM1 expression (C,D) in BeWo cells with suppressed expression of aSMase (A,C) and ceramidase (B,D) compared to transfected controls during FSK-induced differentiation. Data shown represent mean ± SEM of n=3 experiments, compared to controls by two-way ANOVA with Bonferroni test post-hoc.

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Figure 5.12. Basal protein expression of aSMase and ceramidase in FSK-induced differentiation BeWo cells through 7 days of culture. Data shown represent mean ± SEM of n=3 experiments.

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5.2.4. Downstream signaling targets of ceramide regulating its pro-differentiation actions

To identify downstream signaling targets of ceramide responsible for regulating its pro-differentiation role, trophoblasts were treated with inhibitors of common protein targets of ceramide signaling, namely a JNK inhibitor (SP600125), a PP2a inhibitor (okadaic acid) and a PP1 inhibitor (tautomycin) (Pettus et al. 2002). Initially, pilot experiments were undertaken to optimize conditions across a broad concentration range for each inhibitor to ascertain a concentration that could be used without adverse effects of cell viability (Figure 5.13). Cell viability, hCG secretion and PLAP activity were measured in optimization studies to identify the effect of these inhibitors, if any, on biochemical trophoblast differentiation. Trophoblasts were incubated with these inhibitors for up to 5 days, following which cell culture media were collected for hCG secretion assays and cell lysates for PLAP assays. PLAP activity declined significantly in the presence of SP600125 at concentrations 5 µM (~30%), 10 µM (~50%) and 20 µM (~50%) without any detrimental effects on cell viability; however, hCG secretion remained unchanged at these concentrations (Figure 5.13A). Both okadaic acid and tautomycin were toxic at the higher concentrations; however, at concentrations that did not compromise cell viability, neither had any effects on hCG secretion or PLAP activity (Figure 5.13B and C).

The absence of an effect of any of the above mentioned signaling inhibitors on biochemical differentiation as assessed by hCG production argued against these pathways being involved in signaling the effects of ceramide on trophoblast differentiation. Furthermore, incubating trophoblasts with C8 ceramide for 3 h on day 3 of culture resulted in no significant change in phosphorylation state of JNK compared to control untreated cells (Figure 5.14A). Interestingly, while there were no changes in levels of phosphorylation of JNK at site p54, a ~3-fold increase in p46 phosphorylation was observed during trophoblast differentiation in the absence of any other treatment (Figure 5.14B). p46 and p54 are the two isoforms (i.e. with or without a COOH-terminal) that all three JNK genes (JNK1-3) possess.

Ceramide can also activate SAPK proteins and protein phosphatases (PPs), which in turn inhibit MAPKs and classical protein kinases such as PKA and PKB/Akt. We explored the effects of ceramide on Akt phosphorylation by phospho-immunoblotting

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Results: Chapter 5 and found a >80% decrease in Akt phosphorylation after 3 h incubation with C8 ceramide on day 3 of culture (Figure 5.15A). In addition, Akt phosphorylation was upregulated by ~3-fold on day 3 of culture (compared to day 1) when trophoblasts were undergoing differentiation (Figure 5.15B).

Figure 5.13. Effect of SP600125, okadaic acid and tautomycin on hCG secretion, PLAP activity and cell viability by human primary culture of trophoblasts. Cells were treated for up to 5 days with different concentrations of (A) JNK inhibitor SP600125 (0.1, 1, 5, 10 and 20 μM), (B) PP2A inhibitor okadaic acid (1, 5, 10, 50 and 100 nM), (C) PP1 inhibitor tautomycin (1, 10, 50, 100 and 1000 nM), or complete medium (control). Supernatants from day 5 were then assayed for hCG secretion, PLAP activity and cell viability. Data are expressed as mean ± S.E.M. compared to control by two-way ANOVA with Bonferroni test post-hoc.

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Figure 5.14. Phosphorylation of JNK. (A) Immunoblots representing JNK phosphorylation at sites p54 and p46 with or without C8 ceramide. (B) Comparison between phosphorylation state of JNK before (day 1) and during trophoblast differentiation (day 3). Data represent mean ± SEM for n=3 experiments. *P≤ 0.05 compared to control/day1 by two-way ANOVA with Bonferroni test post-hoc.

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Figure 5.15. Phosphorylation of Akt (A) Immunoblots representing Akt phosphorylation after acute exposure to C8 ceramide for 3 h compared to control. (B) Comparison between phosphorylation state of Akt before (day 1) and during trophoblast differentiation (day 3). Data represent mean ± SEM for n=3 experiments. *P≤ 0.05, ***P≤ 0.001 compared to control/day1 by t-test.

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5.3. Discussion

To the best of my knowledge, the current study presents the first experimental evidence of a role of ceramide and its synthesis/metabolic enzymes (aSMase, CERK and ceramidase) in the regulation of biochemical and morphological trophoblast differentiation in the human placenta. These findings revealed changes in endogenous ceramide levels during trophoblast differentiation, with elevated levels coinciding with onset of biochemical differentiation followed by a decline with syncytialization as levels of metabolic enzymes (CERK and ceramidase) increased. We also observed significant increases in total cellular aSMase expression in trophoblasts undergoing differentiation in vitro (day 3-4), coinciding with elevated levels of endogenous ceramide and increased hCG secretion. Four of the five major ceramidase isoforms have shown altered expression levels during differentiation of keratinocytes as mentioned in Chapter 1, the exception being alkaline ceramidase-2 (Houben et al. 2007). These data imply an essential role of ceramidases in cell differentiation and consequently developing of a functional skin barrier (Houben et al. 2007).

Although the major ceramidases present in the human placenta have yet to be characterized, data from different cellular systems suggest that acid ceramidases are the predominant isoforms (Houben et al. 2007). The progressive increase in expression of CERK and ceramidase during trophoblast differentiation and fusion in vitro suggests they may play more of a role in maintenance of the syncytium rather than in the differentiation process. Interestingly, the gradual increase in CERK expression during differentiation is in contrast to findings in the leukemia cell line HL-60, which showed a decrease in CERK expression during macrophage-like cell differentiation (Date et al. 2007). It should, however, be appreciated that the role of these molecules is very cell dependent and reports of opposing actions in different cell types are not uncommon.

Ceramide is a ubiquitous SPL at the heart of a complex web of synthetic and metabolic pathways (Sweeney et al. 1998; Pettus et al. 2002; Rotolo et al. 2005). Distributed primarily in Golgi, ER and plasma membrane, its intracellular location influences its fate and activity (Cremesti et al. 2002; van Blitterswijk et al. 2003). Having been extensively studied with respect to its pro-apoptotic functions (Pettus et al. 2002; Taha et al. 2006; Morales et al. 2007), one of the key stimuli for ceramide generation is activation of early apoptotic

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As discussed in Chapter 3, although initially considered a key component of villous trophoblast differentiation, a recent model proposed the proform and proteolytically active forms of caspase 8 may potentially have different roles. While the former was implied to contribute to early biochemical differentiation, the latter cleaving fodrin at the last steps of fusion (Rote et al.). In agreement with this model, the lack of effect in the present study of ceramide, aSMase and B13 on certain fusion markers was mirrored by a similar lack of effect on activity of caspase 8. In our hands ceramide failed to affect caspase 8 activation or cell viability in primary trophoblasts. The absence of an effect of C8 ceramide on cell fusion, as indicated by E-cadherin immunostaining, may reflect a compartmentalized role of ceramide in the differentiation process independent of syncytial fusion. It is reasonable to question whether these effects are exclusive to short chain ceramides, which have been shown to display different properties to their naturally occurring long chain counterparts due to their hydrophilic nature and ability to easily be taken up by cells (Gangoiti et al. 2010). However, the C8 ceramide data in this study are consistent with the bacterial aSMase administration data as discussed above, which generates release of long chain intracellular ceramide, supporting the validity of the results. Additionally, previous ceramide apoptosis studies in trophoblasts have found little difference in the ability of short (C2) or long (C16) chain ceramides to induce apoptosis (Payne et al. 1999).

Consistent with the hypothesis that these findings represent a causal association between ceramide metabolism and differentiation, exogenous administration of a cell permeable short chain ceramide enhanced biochemical trophoblast differentiation in vitro, as illustrated by increased hCG secretion levels plus a decrease in GCM1 expression. However, the changes in biochemical differentiation were not reflected by altered morphology/amount of syncytialization. Treatment with bacterial aSMase had no significant effect on GCM1 expression or syncytial formation, although it did enhance hCG secretion. Inhibition of acidic ceramidase activity (Delgado et al. 2006) resulted in upregulation of trophoblast differentiation (hCG secretion), but paradoxically down-regulation of syncytial formation. These findings either reflect non-specific effects of the inhibitor, which is highly unlikely considering its reported

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On the other hand, exposure to alkaline and neutral ceramidase inhibitor D-erythro- MAPP failed to induce any effect in differentiation and fusion (Bielawska et al. 2007), supporting the idea that there are distinct signaling pools of ceramide in trophoblasts, and suggesting the acidic isoform of ceramidase is most likely predominant with respect to regulation of trophoblast differentiation.

Another ceramide-synthesizing enzyme studied was CerS which generates ceramide via two pathways (acylation of sphinganine and Sph), although the more predominant pathway is reportedly via sphinganine through the de novo pathway (Soriano 2005).

Lack of an effect on hCG levels by inhibiting CerS using fuminosin B1 was observed in this study, and whether this reflects an absence of the de novo pathway in trophoblast differentiation, a lack of involvement in differentiation or rapid elimination of fuminosin B1 from trophoblasts converted is yet to be determined. Alternatively, it may also be likely that this particular CerS isoform (out of the six in total) targeted by fuminosin B1 is not functionally related to trophoblast differentiation.

The control of hCG secretion (as opposed to hCG expression) is poorly understood, although changes in intracellular cAMP, calcium and PKC activity have been implicated, all of which may be responsive to ceramide modulation (Newhouse et al. 2007). In our studies, modulating ceramide levels acutely after hCG secretion levels declined during biochemical differentiation failed to show any significant effects on hCG secretion, suggesting that the effects of ceramide on hCG secretion were a consequence of changes in biochemical trophoblast differentiation, not more acute effects on hCG production and release. The differences observed with hCG secretion and expression in response to ceramide modulators and ceramidase inhibitor may perhaps be related to the disparate functions of secreted hCG compared to that of its intracellular counterpart as discussed in the Chapter 3.

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Ceramide-activated downstream signaling proteins include stress-activated protein kinases (for example JNK) and protein phosphatases PP1 and PP2A, via which ceramide inhibits pro-growth cellular regulators such as various PKC isoforms, PKB/Akt and Blc2 (Ruvolo 2001; Ruvolo 2003). Inhibiting JNK, PP2A and PP1 in the current study failed to alter levels of trophoblast differentiation, suggesting that they are unlikely to be the potential targets via which ceramide exerts its pro- differentiation role. However, a significant upregulation in JNK phosphorylation at site p46 was observed as cells differentiated. Interestingly, while inhibition and phosphorylation studies showed no effect of ceramide on JNK activation, a marked decline in Akt phosphorylation was observed in response to ceramide. Our findings present novel data of an association between JNK, Akt and trophoblast differentiation, in line with their recently discovered cell differentiation related role in other biological systems (Wagner and Nebreda 2009; Chen et al. 2010). It should be appreciated here that SPL signaling is extremely complex; ceramide may exert its pro-differentiation role via upstream signaling targets, such as atypical PKCζ, PP1, cathepsin D, kinase suppressor of Ras, or double-stranded RNA-dependent protein kinase (PKR), which in turn may inhibit downstream Akt (Ruvolo 2001).

In this study we have employed a well-validated model of in vitro differentiation based on spontaneous fusion of trophoblasts to examine the regulation of the syncytialization process that is central to syncytial formation and regeneration throughout pregnancy. Perturbations in trophoblast differentiation have been associated with faulty syncytial formation observed in pregnancy complications including IUGR and preeclampsia (Brosens et al. 1977; Himmelmann et al. 1996). Recent findings show that IUGR and preeclampsia have distinct pathologies with different trophoblast differentiation profiles (Newhouse et al. 2007). While trophoblast cultures from IUGR placentas show increased trophoblast differentiation and syncytial fusion compared to normal cells, trophoblasts from preeclamptic pregnancies show only a moderate increase in syncytialization and no difference in differentiation (hCG accumulation) (Newhouse et al. 2007). Interestingly, placentas from pregnancies with both IUGR and preeclampsia have been found to exhibit significantly different hormone accumulation and syncytialization profiles compared to those with IUGR only (Newhouse et al. 2007). These data underscore the fact that

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While the choriocarcinoma BeWo cell line has been used in vitro to study various aspects of trophoblast biology, the nature of BeWo differentiation, syncytial formation and protein expression patterns differs from primary trophoblasts {King, 2000 #1062; Bilban, #1235}. In the present study, silencing of aSMase and ceramidase expression was carried out in BeWo cells using siRNA, as transfection of primary trophoblasts using calcium phosphate-based methodology was unsuccessful, although in my hands it has been used successfully to suppress expression of trophoblast transporter proteins by up to 70% (Chapter 4). Regardless, successful silencing of aSMase and ceramidase expression in BeWo cells failed to show any effects on FSK-induced differentiation as assessed by hCG secretion. Importantly, we found significant differences in expression trends of SPL enzymes during differentiation between trophoblast and BeWo cells, with the later cell type showing no changes in expression of these enzymes. Therefore, we suggest that the BeWo model is not particularly representative of trophoblast SPL metabolism and function and that the gene silencing data should be interpreted accordingly.

In conclusion, in this study we have presented evidence for a role of ceramide and its synthesis/metabolic enzymes in regulating biochemical and morphological aspects of trophoblast differentiation. These findings shed light on an important yet poorly understood process that has significant implications for our understanding of placental developmental disorders. Our findings that ceramide is an intracellular regulator able to exert differential effects on functional trophoblast differentiation and syncytialization raise questions regarding the role of ceramide synthesis and metabolism in abnormal pregnancies. Animal studies suggest that disturbance in SPL metabolism may contribute towards pregnancy loss (Mizugishi et al. 2007). Acidic SMase deficiency has been associated with various complications such as atopic dermatitis, caused by a defective permeability barrier (Houben et al. 2007), and Niemann-Pick disease Type A, which results in accumulation of SM in the placenta from the early stages of pregnancy (Schoenfeld et al. 1985). On the other hand, acid ceramidase deficiency (Farber disease) is an inherited lysosomal lipid storage disorder that results in accumulation of ceramide, which has been associated with intrauterine fetal deaths (van Lijnschoten et al. 2000; Houben et al. 2007). These disorders

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CHAPTER 6. REGULATION OF TROPHOBLAST DIFFERENTIATION AND SYNCYTIALIZATION BY SPHINGOSINE AND S1P

6.1. Introduction

SPLs have only recently become well known as key lipid mediators, intra- and extracellular signaling molecules, and cellular regulators with their biological significance now being widely appreciated. Synthesis of bioactive SPLs induced by a variety of stimuli results in the triggering of a range of downstream pathways and targets that mediate various effects on cell function (Bartke and Hannun 2009). The cellular response depends on the nature and the spatial and temporal characteristics of the pathways that are stimulated in response to SPL release (Kitatani et al. 2008). Sph is generated from catabolism of complex SPLs, such as glycosphingolipids, SM and ceramide, in acidic lysosomal compartments from where it moves freely between cellular membranes. Sph is subsequently converted by enzymes such as SPHK and CerS to bioactive metabolites S1P and ceramide, respectively (Riboni et al. 1998; Kitatani et al. 2008; Bartke and Hannun 2009). S1P, also a product of SPL breakdown, is in turn either dephosphorylated to regenerate Sph by S1PP or metabolized to release hexadecenal and ethanolamine phosphate by S1P lyase (Johnson et al. 2003; Bandhuvula and Saba 2007). Recent studies have shown a reduction in extracellular S1P levels following its dephosphorylation by LPP. The resulting Sph regenerated is taken up by cells then rephosphorylated via SPHK1, thereby enhancing intracellular S1P levels (Zhao et al. 2007).

S1P is a potent, pleiotropic extracellular phospholipid messenger that activates a family of cell surface receptors and exerts a wide variety of effects on cell types throughout the body; in many instances its actions antagonize those of ceramide (Smith et al. 2000; Spiegel and Milstien 2003). Genetic defects in metabolic enzymes involved in Sph synthesis/metabolism in acidic late endosomes/lysosomes lead to lysosomal accumulation of the substrate lipids, causing a group of disorders termed as sphingolipidoses, which include Farber’s disease, Niemann-Pick type A and B diseases, and Gaucher disease as briefly mentioned in Chapter 5 (Park and Schuchman 2006; Eliyahu et al. 2007; Kitatani et al. 2008; Bartke and Hannun 2009). SPHK is a

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Results: Chapter 6 highly conserved enzyme important for cell signaling, and disruption of SPHK has been associated with severely defective decidualization, resulting in miscarriages (Mizugishi et al. 2007) (Maceyka et al. 2002; Spiegel and Milstien 2003).

Placental growth and development requires villous trophoblast differentiation and fusion formating a multinucleated syncytium, a unique process that occurs throughout pregnancy (Gauster et al. 2009). Perturbations in trophoblast differentiation have been associated with faulty syncytial formation causing inefficient nutrient transfer across the placenta, which eventually leads to pregnancy complications (Brosens et al. 1977; Himmelmann et al. 1996). S1P receptors are expressed in the human placenta (Johnstone et al. 2005; Hemmings et al. 2006; Hong et al. 2008), as is SPHK1, although little is known of the production and roles of Sph and S1P in normal and abnormal placental physiology. Johnstone et al. in an in vitro study of primary term placental trophoblasts, presented evidence that S1P has an inhibitory effect on trophoblast differentiation, mediated via inhibition of cAMP generation or actions (Johnstone et al. 2005). A role of Sph has also been reported in differentiation studies in leukemia cells; however, there have not been any reports on Sph regulating trophoblast differentiation to date (Shirahama et al. 1997). As mentioned in the previous chapter, regulation of trophoblast differentiation involves ceramide and its primary synthetic and metabolic enzymes, aSMase and acid ceramidase (Singh et al. 2010). Based on these findings, and insight from the “ceramide-S1P rheostat” concept observed in models of cellular apopotosis, I hypothesized that Sph and the activity of key S1P-synthesizing enzyme, SPHK1, are involved in regulation of villous trophoblast differentiation and syncytialization (Okazaki et al. 1989; Hannun and Obeid 2002; Futerman and Hannun 2004).

6.2. Results

6.2.1. Changes in intracellular levels of Sph, secreted S1P and expression of their biosynthesis/metabolic enzymes during spontaneous trophoblast differentiation.

Using immunoblotting analysis of proteins extracted from term villous trophoblasts as they differentiate in culture, I investigated the expression patterns of SPHK1, the key

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Results: Chapter 6 enzyme responsible for phosphorylation of Sph to S1P, and S1PP2, which regulates the reverse of SPHK1 activity. As shown in Figure 6.1A, levels of SPHK1 protein expression were relatively high in undifferentiated CTBs and remained so throughout the early phases of trophoblast differentiation, then drastically declined by ~80% after day 3 of culture and remained low as syncytialization was achieved and the cells became terminally differentiated. I also investigated the expression levels of SPHK2 and S1PP2 during trophoblast differentiation. While SPHK2 expression was undetectable, S1PP2 expression levels remained unaltered as cells differentiated (Figure 6.1B); therefore these enzymes were not investigated further in this particular study.

Levels of intracellular Sph decreased modestly through the differentiation process and had declined by 30% in terminally differentiated cells (Figure 6.2A). Mean intracellular amount of Sph on day 1 was 2330 mg. On the other hand, while intracellular S1P levels were below the limit of detection, concentrations of S1P in conditioned media (containing charcoal-stripped serum) showed a dramatic 10-fold reduction on day 3 of culture, with a rapid increase back to basal levels by day 5 (Figure 6.2B). Collectively, these changes indicate diminished production of Sph and S1P during differentiation in vitro, tying in with reduced expression levels of Sph- metabolizing and S1P- generating enzyme SPHK1.

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Figure 6.1. Expression of SPHK1 and S1PP2 during villous trophoblast differentiation in vitro. Western blotting was used to determine protein expression patterns of SPHK1 (A) and S1PP2 (B) during 7 days in culture during spontaneous villous trophoblast differentiation. Data shown represent mean ± SEM of n=3-4 experiments. **P ≤ 0.01, ***P≤ 0.001 by one-way ANOVA with Bonferroni test post-hoc.

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Figure 6.2. Levels of intracellular Sph and secreted S1P during trophoblast differentiation in vitro. Levels of intracellular Sph (A) and secreted S1P (B) were measured by LC-MS/MS over 7 days in culture during spontaneous villous trophoblast differentiation. Data shown represent mean ± SEM of n=3-4 experiments. *P ≤ 0.05 compared to day 1 by one-way ANOVA with Bonferroni test post-hoc.

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6.2.2. Regulation of trophoblast differentiation by the Sph-S1P pathway.

To investigate the role of the Sph-S1P pathway in trophoblast differentiation, cells were exposed to 10 µM synthetic Sph (Yoon et al. 2009), S1P (Johnstone et al. 2005) and SPHK1 inhibitor (French et al. 2003; Pchejetski et al. 2005) for up to 72 h (Figure 6.3).

Figure 6.3. Illustration of SPLs and enzymes in the Sph-S1P pathway targeted with modulators or SPHK inhibitor to investigate their role in trophoblast differentiation and syncytialization

None of the treatments had any toxic effect on the cells as assessed by MTT assay (Figure 6.4A and B). Indeed, the SPHK1 inhibitor unexpectedly caused a small but significant increase in MTT activity at the 24- and 48-h time points (Figure 6.4C). In addition, these exogenous manipulations failed to exert any changes in caspase 8 activity (Figure 6.4D, E and F).

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Measurement of hCG secretion as an index of biochemical differentiation revealed that all of these treatments inhibited differentiation as shown by reduced hCG secretion at the 48 and/or 72 h time point (~50% in the case of Sph and SPHK1 and ~70% in response to S1P) (Figure 6.5A, B and C). Intracellular levels of hCG were also determined after exposure to these compounds. While SPHK1 inhibition exerted a similar effect on hCG expression as it did on secretion (Figure 6.5F), none of the other treatments affected hCG expression significantly, although the trends were similar to hCG secretion studies (Figure 6.5D, E). Inhibition of SPHK1 resulted in a ~70% downregulation of hCG expression at 48 h and ~60% at 72 h time points (Figure 6.5F). To determine whether any of these particular Sph-S1P pathway compounds had direct effects on hCG secretion that may be independent of their effects on differentiation, trophoblast cultures were exposed to modulators/inhibitors for 24 h on day 4 of culture (i.e. post-differentiation). As shown in bottom panel of Figure 6.5, no significant changes in hCG secretion were observed with Sph (Figure 6.5G), S1P (Figure 6.5H) or SPHK1 inhibitor (Figure 6.5I) compared to controls.

N,N-dimethylsphingosine (DMS; 5 µM), another widely used (but non-selective) inhibitor of SPHK1, was also employed in these studies (Jamal et al. 1991; French et al. 2003). While it significantly downregulated hCG secretion by over 10-fold (Figure 6.6B), DMS treatment had toxic effects on the cells after 48 h decreasing viability by 50% (Figure 6.6A). By 72 h of DMS exposure to cells, viability had decreased by ~70% (Figure 6.6A) due to which further use of this inhibitor was discontinued in these studies. Thus, the DMS-related toxicity may explain the concurrent decrease in hCG production in these experiments.

The mRNA expression of GCM1 was also measured in response to these treatments. GCM1 expression was unaffected by Sph (Figure 6.7A), but showed a transient ~40% decrease in expression at 24 h in response to S1P (Figure 6.7B). Exposure to the SPHK1 inhibitor resulted in a significant downregulation (~70%) in GCM1 expression at 24 h that was maintained up to 72 h of culture (Figure 6.7C). PLAP activity tended to be reduced in response to treatment, although this did not reach statistical significance (Figure 6.7D-F); however; in contrast to the other data and trends, S1P caused an unexpected upregulation in PLAP activity (~40%), but only at the final time point (Figure 6.7E).

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Cell fusion was assessed immunocytochemically using E-cadherin staining to visualize cell-cell boundaries and counting the relative numbers of multinuclear cells (2 or more nuclei per cell) vs. total number of nuclei on day 5 of culture (Figure 6.8A- D). Syncytial formation was investigated following administration of Sph or S1P, or SPHK1 inhibitor (Figure 6.8E). No significant changes in amount of cell fusion was detected over 5 days in culture in response to Sph, whereas S1P resulted in a modest but significant decrease in amount of cell fusion by ~15%. Conversely, inhibition of SPHK1 activity resulted in an upregulation of cell fusion by ~7% (P<0.05).

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Figure 6.4. Effects of Sph, S1P and SPHK1 inhibitor on trophoblast cell viability. Cells were treated with Sph (A), S1P (B) and SPHK1 inhibitor (C) for up to 72 h and viability was determined by MTT assay. Caspase-Glo® 8 Assay was used to study caspase 8 activity in response to Sph (D), S1P (E) and SPHK1 inhibitor (F). Results are expressed as a percentage of MTT reduction and activity of caspase 8, respectively, compared to control conditions for n=3-4 experiments by two-way ANOVA with Bonferroni test post-hoc. **P≤ 0.01.

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Figure 6.5. Regulation of hCG levels by SPL compounds. Shown are the effects of exogenous administration of Sph (A, D, G), S1P (B, E, H) and SPHK1 inhibitor (C, F, I) on hCG secretion and intracellular expression of hCG during differentiation, and post-differentiation hCG secretions levels on day 5 compared to vehicle-treated cells. Data are mean ± SEM of n=3-5 experiments. *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001 compared to controls for each treatment at respective time points by two-way ANOVA with Bonferroni test post-hoc.

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Figure 6.6. Effect of DMS on trophoblasts during differentiation. Cell viability (A) and changes in hCG secretion (B) in response to inhibition of SPHK1 using DMS (5 µM) compared to vehicle-treated cells during differentiation. Data are mean ± SEM of n=3-5 experiments. *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001 compared to control by two-way ANOVA with Bonferroni test post-hoc.

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Figure 6.7. Regulation of biochemical differemtiation markers in response to SPL modulators or enzyme inhibitors. Shown are the effects of exogenous administration of Sph (A, D), S1P (B, E) and SPHK1 inhibitor (C, F) on GCM1 expression and PLAP activity compared to vehicle-treated cells during differentiation. Data are mean ± SEM of n=3-5 experiments. *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001 compared to controls for each treatment at respective time points by two-way ANOVA with Bonferroni test post-hoc.

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Figure 6.8 Changes in morphological differentiation. Immunocytochemistry (A-D) was used to determine the ratio of multinuclear cells (2 or more nuclei per cell) to total number of nuclei, assessed by E-cadherin expression, in response to Sph, S1P and SPHK1 inhibitor (E) compared to untreated cells. Data shown represent mean ± SEM of n=3 experiments. *P ≤ 0.05 compared to controls (one-way ANOVA with Bonferroni test post-hoc); scale bar- 20 µm. Multinuclear cells are indicated by arrows.

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6.2.3. Effects of silencing SPHK1 on trophoblast differentiation

Attempts to silence expression of SPHK1 in primary term trophoblasts using calcium phosphate-based transfection protocol were not successful, similar to aSMase and ceramidase transfection studies mentioned in the previous chapter. Although a 62% downregulation in SPHK1 mRNA expression was observed 24 h after trophoblasts were transfected with StealthTM siRNA, the effect was abolished within the next 24 h (Figure 6.9A). We repeated the study in the BeWo choriocarcinoma cell model to investigate the effects of modification of Sph-S1P pathway on differentiation. Expression of SPHK1 in BeWo cells was suppressed by siRNA-mediated transfection by ~80% (Figure 6.9B). However, this had no significant effect on hCG secretion (Figure 6.9C) or GCM1 expression (Figure 6.9D).

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Figure 6.9. Efficiency of siRNA-mediated transfection in primary trophoblasts compared to immortalized BeWo cells. Administration of StealthTM siRNA using calcium phosphate to transfect cells and suppress expression of SPHK1 in trophoblasts (A) and BeWo cells (B). Effects of SPHK1 suppression on hCG secretion (C) or GCM1 expression (D) in BeWo cells compared to transfected controls during FSK-induced differentiation. Data shown represent mean ± SEM of n=3 experiments. *P ≤ 0.05, **P ≤ 0.01, ***P≤ 0.001 compared to transfected controls by two-way ANOVA with Bonferroni test post-hoc.

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6.2.4. Downstream signaling targets of S1P regulating its anti- differentiation actions.

Results from the previous chapter showed a marked increase in Akt activity occurs during spontaneous trophoblast differentiation (Singh et al. 2010). In light of this finding, and of published evidence that S1P can modulate Akt activity (Van Brocklyn et al. 1998; Gonda et al. 1999; Lee et al. 1999; Malek et al. 2001), I explored the effects of S1P on Akt phosphorylation by phospho-immunoblotting. A ~86% decrease in Akt phosphorylation was observed after 3 h incubation with S1P on day 3 of culture (Figure 6.10).

Figure 6.10. Phosphorylation of Akt in response to S1P treatment. (A) Immunoblots representing total and phosphorylated Akt (Ser473) after acute exposure to S1P for 3 h compared to control. Data represent mean ± SEM for n=3 experiments. *P≤ 0.001 compared to control by two-tailed t-test.

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6.3. Discussion

To the best of our knowledge, the current study presents the first experimental evidence of a role of the bioactive lipid Sph and its metabolizing enzyme SPHK1 in the regulation of villous biochemical and structural trophoblast differentiation in term human placenta. While endogenous Sph levels and SPHK1 expression declined during this process in a time dependant manner in vitro, the end product in this pathway, S1P, could not be detected intracellularly in our cultures. However, secreted S1P levels were detectable and dropped steeply as hCG secretion and ceramide levels were elevated, with a return to basal levels after they had undergone syncytial formation.

Decreased Sph levels may best be explained by the actions of the salvage pathway through which lysosomally-synthesized Sph is translocated via vesicular/non- vesicular routes to the plasma membrane where it becomes available for ceramide synthesis, conversion to S1P and/or efflux into the extracellular milieu; this leads to decreased intracellular levels as differentiation progresses (Herget et al. 2000). It is noteworthy that all Sph is strictly a product of SPL breakdown, contrary to its counterpart dihydrosphingosine that is mostly generated by de novo SPL biosynthesis (Kitatani et al. 2008). Sph generation predominantly relies on the actions of ceramidases which are thus responsible not only for ceramide metabolism but also the synthesis of both Sph and S1P in cells (Mao and Obeid 2008). The salvage pathway encompasses SPLs and glycosphingolipids generated from reacylation of Sph being distributed to plasma membranes and subcellular organelles undergoing turnover with degradation and regeneration. Accordingly, this pathway contributes from 50% to 90% of SPL biosynthesis, underscoring the relevance of SPL breakdown in SPL turnover/biosynthesis and cellular signal transduction (Gillard et al. 1998).

S1P acts via binding to extracellular receptors; therefore, its secretion from the cell is necessary to exert its actions (Alvarez et al. 2007; Hannun and Obeid 2008; Pyne et al. 2009). However, the turnover time for S1P is very rapid with a 15 minute half life, indicative of highly active synthesis and degradation pathways (Venkataraman et al. 2008). Moreover, S1P also acts as a key bioactive molecule via its functions as an intracellular second messenger (Payne et al. 2002). In line with our S1P data, a significant concentration gradient between intra- and extracellular levels of S1P has been previously cited. Studies have shown rapid efflux and externalization of

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Results: Chapter 6 intracellularly synthesized S1P to the outer membrane, where it acts as a extracellular messenger (Caligan et al. 2000; Payne et al. 2002; Berdyshev et al. 2005). Data from the last chapter showed that endogenous levels of ceramide, a S1P precursor, increase during biochemical differentiation and fusion (Singh et al. 2010); my present data showing S1P levels declining during this phase is an example of the well coordinated homeostatic balance during SPL metabolism (Singh et al. 2010). It is of interest to note that cellular levels of ceramide, Sph and S1P differ significantly, with highest levels exhibited by ceramide and S1P presenting the lowest levels. Thus, slight changes in ceramide levels may cause considerable alterations in levels of Sph and S1P (Bartke and Hannun 2009). This makes studying S1P regulation rather challenging due to its rapid turnover rate and dual functions both as an intracellular second messenger and an extracellular ligand for a family of five G-protein-coupled receptors, three of which were recently found to be expressed in the placenta (Hong et al. 2008).

S1P metabolism is catalyzed by S1PP1 and 2, S1P lyase and LPPs (Zhao et al. 2007). While S1PP2 expression remained unchanged during trophoblast differentiation, the activation and/or functional significance of the other S1P metabolizing enzymes during functional and structural trophoblast differentiation is yet to be studied, and may explain the experimentally undetectable levels of intracellular S1P. Alternatively, changes in S1P efflux may explain the drop in media levels of S1P at the time of onset of differentiation. Interestingly, LPPs have been shown to decrease extracellular S1P concentrations by increasing its dephosphorylation to generate Sph which is taken up by cells and converted to S1P by SPHK1 resulting in enhanced intracellular levels (Zhao et al. 2007). However, from our studies, it seems that secreted S1P is the more abundant form in trophoblasts since intracellular S1P levels were undetectable. Further work is required to confirm the role of specific S1P receptors in mediating the effects of extracellular S1P on trophoblast differentiation (Maceyka et al. 2002).

Investigations to identify the precise role of Sph, S1P and SPHK1 activity in villous trophoblast fusion revealed disparities between biochemical differentiation markers and those associated with syncytialization. It is becoming increasingly recognized that at least some aspects of trophoblast differentiation can be dissociated judging by differences between morphological (E-cadherin expression) and biochemical (hCG secretion) differentiation markers (Kao et al. 1988; Al-Nasiry et al. 2006). While Sph

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Results: Chapter 6 caused a decline in hCG production in my current study, it had no significant effect on other biochemical markers (GCM1 expression and PLAP activity) or the amount of cell fusion. SPHK1 activity inhibition decreased hCG production and GCM1 expression; however, it enhanced cell fusion implying SPHK1 activity upregulates trophoblast differentiation while negatively controlling syncytial formation. Discrepancies observed in differentiation regulation in response to exogenously administered Sph and that of accumulated intracellular Sph by inhibiting SPHK1 may be explained by Chigorno et al.’s data showing selective fates of free sphingosine (Chigorno et al. 2005). While the majority of endogenous Sph is recycled, exogenous Sph taken up by cells is directly transported to the lysosomes largely to be metabolized to water and small breakdown products, presumably by first being phosphorylated to S1P and then converted to ethanolamine and palmitaldehyde (Chigorno et al. 2005).

On the other hand, while the current S1P results agree with findings of Johnstone et al. with respect to inhibition of hCG production, I observed differences in the effect of S1P on PLAP activity and cell fusion (Johnstone et al. 2005). Their study showed S1P negatively regulating hCG secretion without modulating fusion (Johnstone et al. 2005), whereas we observed parallels between the effect of S1P on hCG secretion and cell fusion rates, with an attenuation in both in the presence of S1P. These discrepancies may be due to differences in culture conditions and choice of basal medium used. It is also noteworthy that the anti-differentiation role of these bioactive compounds is completely opposite to their role observed in epidermal cell differentiation (Wakita et al. 1994; Johnstone et al. 2005; Sun et al. 2008). It is becoming increasingly apparent that the roles of these molecules are very cell dependent with apparently opposing actions often seen in different studies using different cells.

The SPHK1 isoform has been more intensely researched than SPHK2 (Meacci et al. 2008); the limited research on SPHK2 indicates the two isoforms can have different subcellular localizations and may also exert opposition actions (Spiegel and Milstien 2003). While several studies have reported on the stimulatory effects of SPHK1 activity in cell proliferation and survival, there are some that attribute cell cycle arrest to SPHK2 (Igarashi et al. 2003). My preliminary studies failed to detect any SPHK2 expression during trophoblast differentiation. Although a number of studies have

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Results: Chapter 6 suggested that S1P/SPHK1 act in a growth promoting fashion, others have observed an anti-proliferation/pro-differentiation function of these compounds (Olivera et al. 1999; Pyne and Pyne 2000; Edsall et al. 2001; Donati et al. 2005; Meacci et al. 2008). The varied biological responses elicited by S1P may be due to differential expression of the S1P receptors in the different cell types, which are known to be coupled to divergent biological effects (Ishii et al. 2004). To some extent, the effects observed in the present study of Sph and S1P on trophoblast differentiation conflict with those of the SPHK1 inhibitor, a highly specific inhibitor of SPHK1 (French et al. 2003). I observed increased cell fusion and decreased GCM1 expression with the SPHK1 inhibitor, both consistent with the predicted effects of a decrease in the inhibitory effects of S1P. However, reduced hCG production was also detected, which is opposite to the expected response to inhibition of S1P production. This anomaly might be explained by direct effects of intracellular S1P on the regulation of hCG expression or release, independent of any effects of differentiation. It may also be possible that SPHK1 exerts non-lipid kinase actions which, when inhibited, might result in reduced hCG production (Pyne et al. 2009).

The opposing cellular effects of ceramide and S1P identified in this project during villous trophoblast differentiation mirror those observed in apoptosis and cell growth, highlighting the complex nature of SPLs regulation and actions (Maceyka et al. 2002). The divergent effects might be due to multiple stimuli activating these precursors either directly or indirectly, and/or the complex relationship between different lipids generated. Moreover, if both substrate and product are signaling lipids it may contribute towards diverse biological effects (Futerman and Hannun 2004). The opposing actions of S1P and ceramide presented in the current study during trophoblast differentiation suggest that a ceramide-S1P rheostat operates during placental development to balance their divergent effects on differentiation and fusion.

Extracellular S1P binds to its receptors and triggers activation of a wide range of downstream signaling proteins including PI3K, Akt, ERK, p38MAPK, small GTPases Rac and Rho, PLD and intracellular calcium mobilization, resulting in numerous biological responses (Van Brocklyn et al. 1998; Gonda et al. 1999; Lee et al. 1999; Malek et al. 2001). Johnstone et al. proposed that S1P mediated its anti-differentiation role in trophoblasts by inhibiting cAMP production (Johnstone et al. 2005). Interestingly, I observed a dramatic decline in Akt phosphorylation in the presence of

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S1P. I have shown that Akt phosphorylation is markedly increased with trophoblast differentiation in the previous chapter (Singh et al. 2010). This is concordant with studies that show Akt inhibits p38MAPK, which is downregulated during trophoblast differentiation (Daoud et al. 2005). Therefore, it is plausible that the negative effects exerted by S1P on trophoblast differentiation and syncytialization are via the Akt pathway.

In conclusion, I have identified a novel role for Sph/S1P as negative regulators of biochemical and morphological trophoblast differentiation. I reveal that Sph plays a significant role in biochemical trophoblast differentiation without altering syncytial formation, whereas SPHK1 shows divergent outcomes on biochemical differentiation and cell fusion. My results present an entirely new perspective for exploring the potential of SPL compounds, in particular the Sph-S1P pathway, in placental development.

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CHAPTER 7. CONCLUSIONS AND FUTURE PERSPECTIVES

This thesis has introduced a potentially new dimension in understanding the regulation of human trophoblast differentiation and syncytialization. A significant role of bioactive SPLs in regulating these processes has been identified, with compounds from various metabolic pathways exercising divergent effects. Members of the SMase pathway have been shown to be primarily involved with functional differentiation of villous trophoblasts, while the Sph-S1P pathway is associated with both functional and structural trophoblast differentiation (summarized in Figure 7.1). On the other hand, from the current findings the involvement of the de novo pathway in trophoblast differentiation and syncytialization remains largely elusive, and would need to be studied in much greater detail to understand its role, if any, during placentation.

Figure 7.1. Role of SPLs in trophoblast differentiation and syncytialization. While most of the sphingomyelinase pathway is predominantly associated biochemical trophoblast differentiation, the Sph-S1P pathway regulates both biochemical and morphological aspects of trophoblast differentiation and fusion.

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Although trophoblast differentiation has been actively investigated over the past few decades, various contradicting theories describing the fundamental nature of biochemical and morphological trophoblast differentiation exist. In accordance with recent studies, the current data shows that at least some aspects of trophoblast differentiation can be dissociated as witnessed by differential effects of SPLs on morphological and biochemical differentiation (Kao et al. 1988; Johnstone et al. 2005; Al-Nasiry et al. 2006; Rote et al. 2010). This study highlights the possibility of differential functions of different pools of hCG (secreted vs. intracellular) based on their varied trends as trophoblasts differentiated, aggregated and fused and their differing responses to various treatments. Knowledge of the differences in hCG secretion patterns between primary trophoblasts and BeWo cells raises caution when using these choriocarcinoma cells to simulate primary cell culture models. Although choriocarcinoma-derived BeWo cells have been extensively used in vitro to study villous trophoblast differentiation, it should be appreciated that these cells are transformed and their differentiation, syncytial formation and protein expression patterns differ from primary trophoblasts.

Unlike cells undergoing apoptosis, differentiating trophoblasts lose membrane asymmetry only transiently during cell fusion without terminal progression of the apoptotic cascade. The lack of correlation between changes in caspase activity (early and effector) and trophoblast differention in the present study questions their proposed roles in syncytializaion (Yusuf et al. 2002; Guilbert et al. 2010). It could be postulated that the well defined ceramide-S1P rheostat prevents trophoblasts from undergoing cell death during caspase activation and reorganization of plasma membrane components (hallmarks of apoptosis) as cells undergo differentiation and fusion. Several studies have shown an association between SPLs and the caspase cascade (Sweeney et al. 1998; Pettus et al. 2002). In trophoblasts, ceramide and aSMase have been shown to display pro-apoptotic characteristics in an EGF-inhibitable manner (Payne et al. 1999). However, in the current studies, neither ceramide nor S1P caused any significant changes on caspase 8 activation. It is possible that the different results reflect different cell types and signaling mechanisms. A plausible explanation for the maintenance of homeostasis during trophoboblast differentiation, aggregation and fusion by the ceramide-S1P rheostat is by operating to balance their divergent effects

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Conclusions and Future Perspectives on biochemical differentiation and syncytial formation, instead of via their apoptotic functions.

Currently, well established downstream signaling targets that mediate syncytialization include PKA, ERK1/2 and p38 mitogen-activated protein kinases (Daoud et al. 2005) (Keryer et al. 1998). This thesis presents novel data regarding PKB/Akt and JNK pathways and their association with key processes essential for placental development. Moreover, changes in Akt phosphorylation were detected in response to ceramide and S1P, suggesting that these SPL compounds may be regulating trophoblast differentiation via a potentially novel mechanism. It should be noted that although ceramide and S1P exert differential actions on differentiation they actually both inhibited Akt phosphorylation, consistent with an anti-differentiation event. SPLs have multiple subcellular localizations with distinct functions; moreover, it could also be possible that different pools of Akt also exist, which mediate the differential downstream effects of these SPLs. Alternatively, the inhibitory effect of exogenous ceramide on Akt could be totally independent to its pro-differentiation role. In addition, while Akt is a known inhibitor of JNK, an upregulation in Akt phosphorylation would expect to downregulate JNK phosphorylation; however, this was not observed in the current study. It is likely that the inhibitory effects of Akt on JNK may be cell-specific, although further studies would be required to confirm these speculations. It should also be appreciated that only one of the three Akt isoforms (Akt-Ser473) has been studied in this project, and it may or may not be representative of the other two Akt isoforms. The effects of these SPLs on the other two Akt isoforms are yet to be elucidated.

This study did not support a role for BCRP in trophoblast differentiation in the human placenta in disagreement with an earlier finding using BeWo cells, which may largely be contributed to the differences in differentiation properties between the two cell types, as aforementioned (Evseenko et al. 2007c). Moreover, the current results revealed the lack of any association between SPLs, in particular ceramide and sphingosine-1-phosphate, and BCRP during trophoblast differentiation and syncytialization. Although an upregulation in ceramide levels was detected in BCRP- silenced cells, it occurred prior to a significant decline in BCRP protein expression was observed and the increase was only transient. The increase in ceramide accumulation may be in response to a compensatory mechanism with BCRP

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Conclusions and Future Perspectives expression being suppressed in a time-dependent manner. However, the expression of ceramide synthesis/metabolic enzymes remained unaltered in transfection studies; therefore, this data should be interpreted with caution.

As mentioned earlier, SPLs are primarily found in the external leaflet of the plasma membrane with varying affinities for other lipids resulting in the generation of sub- microscopic microdomains due to non-random distribution of these compounds (Sietsma et al. 2001) (Cremesti et al. 2002). For example, while SM binds tightly with cholesterol, ceramide has low affinity for this sterol resulting in segregation and formation of exclusive ceramide-enriched microdomains (Kolesnick et al. 2000). These ceramide-enriched microdomains facilitate the lamellar-hexagonal transition of lipids, which destabilize membrane lipids resulting in efflux, fusion or budding of vesicles (Cremesti et al. 2002; Blitterswijk et al. 2003).

Synthesis of ceramide via the SMase pathway is known to regulate lipid microdomain formation (Gulbins and Kolesnick 2003). An increase in ceramide levels is detected in plasma membrane protein caveoli-dominant microdomains in response to TNF stimulation, with a subsequent decrease in SM. TNF stimulation leads to the extrusion of nSMase from these rafts, thus indicating that aSMase may be the key SMase regulating SM hydrolysis and ceramide generation following clustering of TNF receptors in the plasma membrane (Veldman et al. 2001). Generation of ceramide leads to fusion of microdomains to patches, which then coalesce to form platforms

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It is widely established that lipid microdomains play a crucial role in receptor- mediated downstream signaling events in response to ceramide (Cremesti et al. 2002). Transbilayer movement of ceramide synthesized on the extracellular leaflet of the plasma membrane to the inner leaflet, by exogenously added SM, has been shown where it gets phosphorylated by CERK to produce ceramide-1-phosphate. The translocation may disrupt lipid asymmetry leading to the translocated form of ceramide being less associated with its structural role and more of a second messenger.

These findings suggest this translocation may be spontaneous and independent of transporter proteins, and perhaps shed light on an intrinsic property of ceramide enabling ceramide signaling (Mitsutake and Igarashi 2007). Considering the current study explored the role of ceramide as a second messenger in regulating functional trophoblast differentiation, its structural role as a membrane microdomain component can not be ignored and is worth investigating in further detail.

Although Johnstone et al. identified expression of S1P1 to S1P3 receptors in trophoblasts, and showed S1P-mediated regulation of trophoblast differentiation via

Gi-protein coupled receptors, the precise S1P receptor involved in mediating S1P’s functions remains unknown. They also showed a decline in cAMP levels as S1P inhibited trophoblast differentiation. It remains unknown whether S1P mediates its effects via an S1P receptor in the present study, if so, then which one and whether its downstream effects via Akt signaling pathway are realted to cAMP. Due to time constraints, these unananswered questions were unable to be addressed and further studies are required to identify the mechanism involved with S1P’s inhibitory effects of trophoblast differentiation and syncytialization.

In conclusion, this study highlights the complexity of the pathways that regulate the trophoblast differentiation process. Different agents (e.g. ceramide) can regulate functional aspects of differentiation independent to trophoblast fusion, while regulators of fusion can have no or even opposite effects on biochemical differentiation. Pregnancy complications such as preeclampsia, where a moderate increase in syncytialization is

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Conclusions and Future Perspectives observed with no effect on biochemical differentiation, may be the end result of perturbations in one or more of these pathways. The findings of this project are of high relevance considering an increasing number of disorders are being linked to disturbed SPL metabolism (Mizugishi et al. 2007). Recent studies have reported defective decidualization causing miscarriage, preeclampsia-associated umbilical cord artery/vein ‘early ageing’ and hypertension being traced back to SPLs (Romanowicz et al. 2010). Acidic SMase deficiency has been associated with various complications such as atopic dermatitis, caused by a defective permeability barrier (Houben et al. 2007), and Niemann-Pick disease Type A, which results in accumulation of SM in the placenta from the early stages of pregnancy (Schoenfeld et al. 1985). On the other hand, acid ceramidase deficiency (Farber disease) is an inherited lysosomal lipid storage disorder that results in accumulation of ceramide, which has been associated with intrauterine fetal deaths (van Lijnschoten et al. 2000; Houben et al. 2007). These disorders emphasize the potential biological significance of SPL metabolism abnormalities in placentation and pregnancy, and highlight several areas for further research.

Further clarification of the precise downstream mechanisms regulating the differentiation-related effects of SPL compounds should be forthcoming in the near future. It is essential the spatio-temporal metabolism of the bioactive lipids be investigated during future studies to better understand the relevance of their timely interactions exert on trophoblast differentiation and syncytialization. Spatial separation of enzymes allows differential activation of these distinct pathways associated with ceramide generation. However, it remains unclear how signaling pools of ceramide are separated from metabolic pools within a cell. Obviously, compartmentalization is an essential aspect for ceramide function (Hannun and Obeid 2002). Metabolism of SPLs in response to extracellular agents is highly complex and can be specific to a cell type, or regulated by signals produced by allosteric mechanisms, post-translational modifications or enzyme expression patterns (Futerman and Hannun 2004). Accordingly, cellular ceramide homeostasis is governed by which ceramide-generating pathway is activated and its spatial-temporal properties (Kitatani et al. 2008). Being metabolically juxtaposed, the regulation of the metabolism of these lipid mediators is of utmost importance in determining cell fate (Taha et al. 2006). Gaining more insight into the different subcellular localizations of

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Conclusions and Future Perspectives these compounds as trophoblasts differentiate and fuse will enhance appreciation of the importance of SPLs during pivotal stages of placental development.

It should be appreciated and strongly emphasized when interpreting current data that the existing knowledge on regulation of SPL pathways in the placenta is still in its infancy and there are several other enzymes and metabolites involved in these pathways that are yet to be studied. These may provide further insight into the functioning/regulation of these pathways and their role in villous trophoblast and syncytial formation by providing a more ‘complete picture’. It will also essentially shed further light on their role in various lipid storage disorders and pathologies during pregnancy, while opening up new avenues for targeted intervention.

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