The Pennsylvania State University

The Graduate School

Intercollege Graduate Degree Program in Genetics

REQUIREMENT OF THE DYNLRB FAMILY LIGHT CHAINS

IN TRANSFORMING GROWTH FACTOR BETA SIGNALING

A Thesis in

Genetics

by

Guofeng Gao

 2007 Guofeng Gao

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

May 2007 The thesis of Guofeng Gao was reviewed and approved* by the following:

Sarah K. Bronson Associate Professor, Department of Cellular and Molecular Physiology Thesis Advisor Chair of Committee Co-Chair, Intercollege Graduate Degree Program in Genetics

Keith C. Cheng Associate Professor, Department of Pathology

Mark Kester Distinguished Professor, Department of Pharmacology

Jiyue Zhu Associate Professor, Department of Cellular and Molecular Physiology

*Signatures are on file in the Graduate School iii ABSTRACT

Transforming growth factor β (TGFβ) is the prototype for a superfamily of related members. TGFβ family signaling controls various fundamental cellular functions, including cell proliferation and migration. Alterations in the TGFβ signaling pathways have been implicated in a vast array of human cancers and other diseases as well.

Despite advances in our understanding of TGFβ signaling transduction, the mechanism of the multifunctional TGFβ signaling is not completely clear yet. Therefore, further studies are required for deeper understanding of its diverse biological responses. DYNLRB1 was identified in the laboratory through a screen for TGFβ receptor-interacting , and it is also a dynein light chain. Dynein is molecular motor that plays many important functions in the cell. I hypothesized that DYNLRB family dynein light chains may play important functions in TGFβ signaling. In this thesis, we investigated the regulation of the function of DYNLRB dynein light chains by TGFβ and their role in TGFβ signaling in mammalian cells and in primary zebrafish (Danio rerio) ovarian follicle cells.

The work in Chapter 2 aimed to determine the involvement of DYNLRB1 in

TGFβ signaling and characterizing the function of DYNLRB1. The results from this chapter have demonstrated that the phosphorylation of DYNLRB1 on serine residues is stimulated by TGFβ in Cos-1 cells and requires TβRII. It is further demonstrated that

DYNLRB1 expression knockdown significantly impaired TGFβ-induced fibronectin expression in MDCK cells. TGFβ-induced DYNLRB1phosphorylation has been shown to be responsible for its recruitment to the dynein motor complex. Therefore, these results indicate a potential role for DYNLRB1 as a TGFβ signaling intermediate, and its iv requirement in TGFβ induction of fibronectin (a major component of the extracellular matrix), which plays important roles in cell adhesion, migration and differentiation.

The experiments in Chapter 3 was designed to test the hypothesis that DYNLRB2 might be involved in Smad3-dependent TGFβ signaling. Human DYNLRB2 is 77% identical to human DYNLRB1. Results in Chapter 3 have demonstrated that TGFβ induction of SBE2-Luc, plasminogen activator inhibitor-1 expression in HaCaT cells and of Smad7-Luc in Hep3B cells, is significantly impaired, after blocking endogenous

DYNLRB2 expression by siRNA. It is known that the induction of these is

Smad3-dependent signaling event. However, similar blocking DYNLRB2 expression does not inhibit the induction of ARE-Lux by TGFβ, which has been demonstrated to be

Smad2-dependent signaling event. Therefore, these results suggest that DYNLRB2 is specifically required in Smad3-dependent signaling. Further, it is demonstrated that

TGFβ-stimulated preferential interaction between DYNLRB2 and Smad3 may be the underlying mechanism for the requirement of DYNLRB2 in Smad3-dependent TGFβ signaling. In addition, results have shown that TGFβ stimulated a rapid recruitment of the DYNLRB2 to the dynein complex, and the TGFβ-induced phosphorylation of

DYNLRB2 is responsible for this TGFβ stimulated recruitment. Collectively, results in this Chapter have demonstrated for the first time that DYNLRB2 is required for Smad3- dependent TGFβ signaling.

The experiments in Chapter 4 tested our hypothesis that in zebrafish ovarian follicle cells the function of zDYNLRB might be regulated by TGFβ, and zDYNLRB might play an important role in TGFβ signaling in such cells. It is shown that zDYNLRB v is rapidly phosphorylated after TGFβ stimulation, for which the TβRII is required. In addition, it is shown that the phosphorylation of zDYNLRB facilitates its rapid recruitment to the dynein complex, which is stimulated by TGFβ. Knockdown experiments in zOFCs by morpholino have demonstrated that zDYNLRB was required for TGFβ induction of TRE-Luc, 3TP-Lux and ARE-Lux. Thus, the results suggest a potential role for zDYNLRB in TGFβ signaling in zebrafish ovarian follicle cells.

Collectively, the experiments in this thesis demonstrated for the first time a requirement of DYNLRB2 dynein light chain in Smad3-dependent TGFβ signaling in mammalian cells, and a requirement of DYNLRB1 as a potential TGFβ signaling component in TGFβ induction of fibronectin, respectively, as well as potential role for zDYNLRB in TGFβ signaling in zebrafish ovarian follicle cells. However, the physiological significance of their functions needs to be addressed in vivo in the future. vi TABLE OF CONTENTS

List of figures...... x

List of tables ...... xiv

List of abbreviation...... xiv

Acknowledgements...... xix

1. Chapter 1. Literature review…..……………………………..……………… .1

1.1 Introduction…………………………………………………….…………….1 1.2 TGFβ signaling pathways……………………………………….…………...3 1.2.1 The Smad pathway……………………………………….. ..…………..6 1.2.1.1 R-Smads…………………………………………………….………... 7 1.2.1.2 Co-Smads………………………………………………….………...... 7 1.2.1.3 I-Smads…………………………………………………….…………..8 1.2.2 The MAPK pathways………………………………………….………...9 1.2.3 Other pathways……………………………………………….……...…11 1.3 Role of TGFβ signaling in cancer and development……………….………..12 1.3.1 Role in cancer………………………………………………….…..…...12 1.3.2 Role in development…………………………………………….……...18 1.3.3 Role in ovarian follicle development…………………………….……..21 1.4 Regulation of TGFβ signaling………………………………………….…….24 1.4.1 Regulation at Ligand production level…………………………….…….25 1.4.2 Regulation of Ligand Processing and Activation…………………….…26 1.4.2.1 Regulation of Ligand availability for Processing………………….…..27 1.4.2.2 Regulation of Ligand Processing……………………………………....29 1.4.2.3 Regulation of TGFβ Ligand activation…………………………….…..29 1.4.2.4 Regulation of the availability of active mature TGFβ ligand……….....31 1.4.3 Regulation of TGFβ receptor level and activity……………………..….32 vii 1.4.3.1 Enhancing effects of TGFβ co-receptors……………………………...32 1.4.3.2 Negatively regulating the activity of TGFβ receptors………………....33 1.4.3.2.1 Physical Blockade of TGFβ receptors……………………...33 1.4.3.2.2 Enzymatically regulating the activity of TGFβ receptors……………………………………...…35 1.4.3.3 Regulating the level of TGFβ receptors………….………..….36 1.4.4 Intracellular regulation…………………………………………………...37 1.4.4.1 Regulating the Smad2/3 activity by phosphorylation…………………..37 1.4.4.2 Terminating the Smad2/3 signaling by dephosphorylation…………….39 1.4.4.3 Terminating the Smad2/3 signaling by irreversible degradation……………………….…………..…….40 1.4.4.4 Regulating the level and activity of the Co-Smad, Smad4……………...41 1.4.4.5 Regulating the subcellular compartmentalization of Smad2/3………….43 1.4.5 Transcriptional regulation of target expression in the nucleus…………………………………………..…….48 2. Chapter 2. Requirement of DYNLRB1 for TGFβ-mediated induction of fibronectin………………………………….53 2.1 Introduction…………………………………………………………………….54 2.2 Materials and methods…………………………………………………………56 2.3 Results………………………………………………………………………….61 2.3.1 TGFβ stimulates DYNLRB1 phosphorylation, which occurs on serine residues……………………………61 2.3.2 DYNLRB1 detection by rabbit polyclonal DYNLRB1 anti-serum……….65 2.3.3 DYNLRB1 has a short half-life and DYNLRB1 specific siRNAs knock down its expression in MDCK cells………...68 2.3.4 DYNLRB1 knockdown reduced fibronectin induction by TGFβ in MDCK cells…………………………………...70 2.3.5 TGFβ stimulates the interaction between DYNLRB1 and dynein motor, and TβRII is required for this interaction…………...72 2.4 Discussion……………………………………………………………………...74 viii 3. Chapter 3. Requirement of DYNLRB2 in Smad3-dependent TGFβ signaling………78 3.1 Introduction……………………………………………………………….……79 3.2 Materials and methods………………………………………………………….81 3.3 Results…………………………………………………………………………..87 3.3.1 DYNLRB2 siRNAs specifically block DYNLRN2 expression…………...87 3.3.2 DYNLRB2 knockdown significantly inhibits TGFβ induction of Smad3-dependent transcriptional activation of SBE2-Luc and Smad7-Luc……………………...90 3.3.3 DYNLRB2 knockdown significantly interferes with TGFβ induced PAI-1 gene expression……………………..…94 3.3.4 DYNLRB2 is in early endosomes with Smad3 after TGFβ stimulation…..98 3.3.5 TGFβ induces a preferential interaction between DYNLRB2 and Smad3…100 3.3.6 TGFβ stimulates the recruitment of DYNLRB2 to the dynein motor……104 3.3.7 DYNLRB2 is phosphorylated after TGFβ stimulation and TβRII is required for this phosphorylation…………………..108 3.3.8 DYNLRB2 phosphorylation mediates its rapid recruitment to the dynein motor, which is stimulated by TGFβ………………..114 3.4 Discussion…………………………………………………………………..…114 4. Chapter 4. Requirement of zebrafi sh dynein light chain zDYNLRB in TGFβ signaling i n zebrafish ovarian follicle cells………….122 4.1 Introduction………………………………………………………………...…123 4.2 Materials and methods……………………………………………………...…127 4.3 Results…………………………………………………………………………131 4.3.1 Cloning and expression detection of zDYNLRB…………………………131

4.3.2 Primary zebrafish ovarian follicle cells (OFCs) are TGFβ responsive…...135

4.3.3 zDYNLRB is phosphorylated after TGFβ receptor activation……….…..137

4.3.4 TGFβ stimulates the interaction between zDYNLRB and DIC,

which requires the TβRII kinase……………………………….140 ix 4.3.5 zDYNLRB specific morpholinos knock down zDYNLRB expression…..148

4.3.6 zDYNLRB knockdown interferes with some TGFβ-induced transcription activation………………………………………..149 4.4 Discussion……………………………………………………………..………157 5. Chapter 5. Overall discussion and future directions………………………………....165 References…………...………………………………………………………………….179 Appendices……………...………………………………………………………………209 Appendix A. Blocking dDYNLRB2 partially impairs TGFβ-mediated DNA synthesis inhibition in MDCK cells with high passage numbers (25-30), but not in MDCK cells with low passage numbers (<15)………………………….………...…210 Appendix B. dDYNLRB2 siRNA specifically knockdown exogenous dDYNLRB2 protein expression…………………...…211 Appendix C. dDYNLRB2 siRNA specifically knockdown endogenous dDYNLRB2 mRNA expression…………..……...…212 Appendix D. dDYNLRB1 siRNA specifically knockdown endogenous dDYNLRB1 protein expression……………..….…..213 Appendix E. Nonspecific development arrest of 24 h and 48 h embryos injected with the ATG MO……….…..…214 x LIST OF FIGURES

Figure Page

1 Model for TGFβ regulation of transcription through Smad signaling proteins...4

2 TβR internalization by clathrin- and lipid-raft-mediated endocytosis………....45

3 Comparison of Smad2 and Smad3……………………………………………..50

4 DYNLRB1 is phosphorylated upon activation of TGFβ receptors…………….64

5 Activation of the TGFβ receptors results in phosphorylation of DYNLRB1

primarily on serine residues….………………………………………...66

6 Specificity assessment of a rabbit polyclonal

antiserum for detection of DYNLRB1 protein expression…………….67

7 Determination of DYNLRB1 protein half-life…………………………………69

8 The siRNA blockade of DYNLRB1 expression reduces TGFβ

induction of fibronectin expression.……………………………71

9 Phosphorylation of DYNLRB1 is required for

recruitment of DYNLRB1 to the DIC…………………………73

10A dDYNLRB2 siRNA specifically knockdown exogenous

dDYNLRB2 protein expression………………………….88

10B hDYNLRB2 siRNA specifically knockdown endogenous

hDYNLRB2 mRNA expression………………………….89

10C siRNA blockade of endogenous hDYNLRB2 expression

results in significant inhibition of Smad3-dependent

SBE2-Luc activation in HaCaT cells……………………...92

10D siRNA blockade of endogenous hDYNLRB2 expression has xi no effect on inhibition of Smad2-dependent

ARE-Lux activation in HaCaT cells………………...…93

10E siRNA blockade of endogenous hDYNLRB2 expression

inhibits TGFβ-mediated transcriptional activation of

human the Smad7 promoter in Hep3B cells……………….....95

11 siRNA blockade of endogenous hDYNLRB2 inhibits TGFβ-mediated induction

of endogenous PAI-1 gene expression……………………...97

12 hDYNLRB2 is present in EEA1-enriched early endosomes

together with TβRII and Smad3 after TGFβ treatment……….…..99

13A hDYNLRB2 interacts preferentially with Smad3

in IP/blot analyses in 293T cells……………………………….....101

13B hDYNLRB2 interacts preferentially with Smad3

in LUMIER analyses in IEC4-1 cells …………………….103

14A TGFβ stimulates the recruitment of hDYNLRB2 to DIC

in 293T cells……………………………………………………...105

14B TGFβ stimulates the recruitment of hDYNLRB2 to DIC

in HaCaT cells………………………………………………………..107

15 DYNLRB2 interacts with the TβRII in 293T cells…………………………..109

16A hDYNLRB2 is phosphorylated upon activation of TGFβ receptors.………...111

16B TGFβ stimulated hDYNLRB2 phosphorylation in both

Mv1Lu epithelial cells and in R1B cells, but not in DR26 cells …….113

17 The interaction between DIC and hDYNLRB2 requires

TβRII kinase activity…………………………………………………..115

18 Amino acid sequence alignment of hDYNLRB1, hDYNLRB2 xii and zDYNLRB………………………………………………..133

19 Detection of zebrafish DYNLRB……………..……………………………..134

20 zOFCs are responsive to TGFβ treatment……………………….………….136

21A zDYNLRB is phosphorylated upon activation of TβRs in 293T

cells and this phosphorylation is blocked by TGFβ KNRII…………139

21B zDYNLRB is phosphorylated in zOFCs upon TGFβ treatment

and KNRII significantly decreased this phosphorylation…………141

22A TGFβ stimulates the recruitment of zDYNLRB to the DIC

in Mv1Lu cells……………………………………………………143

22B The recruitment of endogenous zDYNLRB to the DIC

in zOFCs is stimulated by TGFβ…………………………………..144

22C The interaction between zDYNLRB and the DIC in zOFCs is

confirmed by opposite direction IP/blot analyese …………...146

22D A functional TβRII is required for the TGFβ induction

of the DIC-zDYNLRB interaction……………………………147

23A The zDYNLRB MOs specifically knock down

exogenous zDYNLRB expression in 293T cells………………150

23B The zDYNLRB MOs specifically knock down

endogenous zDYNLRB expression in zOFCs………………....151

24A MOs knocking down zDYNLRB expression markedly

inhibited TRE-Luc induction by TGFβ……………………..…153

24B Blocking zDYNLRB expression also inhibited

3TP-Lux induction by TGFβ…………………………………...154

24C zDYNLRB expression knocking down did not interfere with xiii phTG5-Lux transcriptional regulation……………………….…156

24D Blocking zDYNLRB expression significantly repressed

ARE-Lux induction by TGFβ……………………………….…..158

25 Hypothetical model………………………………………………………166-167 xiv LIST OF TABLES

Table Page

1 Knockout mouse models of TGFβ family signaling proteins……………………...16-17

2 Comparision of DYNLRB1 to some other DYNLRB/robl/LC7 family members…...62

3 TGFβ family signaling pathway component mutants

with ovary development defects………………161 xv LIST OF ABBREVIATIONS

α alpha

AP-1 activator protein 1

ARE activin response element b bases

β beta

BMP bone morphogenic protein

CamKII Calcium-calmodulin-dependent protein kinase II cdk cyclin-dependent kinase

Co-Smad common Smad cPML cytoplasmic form of the ProMyelocytic Leukemia tumor

suppressor protein

Dab2 Disabled 2

°C degrees Celsius

DIC dynein intermediate chain

CREB cyclic AMP response element binding protein

DLC dynein light chain

DNA deoxyribonucleic acid

ECM extracellular matrix

EGF epidermal growth factor

ELF Embryonic Liver Fodrin

EMT epithelial-to-mesenchymal transdifferentiation

ERK extracellular signal-related kinase xvi γ gamma

GADD34 growth arrest and DNA-damage-inducible 34

GDF growth and differentiation factor h hour

HCCC human colon carcinoma cells

HC heavy chain

Hrs Hepatocyte growth factor-Regulated tyrosine kinase Substrate

IC intermediate chain

IP Immunoprecipitation

I-Smad inhibitory Smad

JNK c-jun-N-terminal kinase

KNRII kinase deficient TβRII

LC light chain

LIC light-intermediate chain

LTBP latent TGF-beta binding proteins

LUMIER luminescence-based mammalian interaction mapping

MAPK mitogen-activated protein kinase

MEK mitogen-activated protein kinase min minute mRNA messenger ribonucleic acid

MT1-MMP membrane type 1-matrix metalloprotease

µg microgram

µl microliter xvii ml millililiter

µM micromoles per liter

MO Morpholino phosphorodiamidate oligonucleotide

MT ng nanogram pmol picomole

PAI-1 plasminogen activator inhibitor –1

Pak-1 p21-actvivated kinase

PCR polymerase chain reaction

PI3K phosphatidylinositol-3k-kinase

PI3P phosphatidylinositol 3-phosphate

PIASy protein inhibitor of activated STATy

PKA protein kinase A

PKC protein kinase C

PP1 protein phosphatase 1

PP2A protein phosphatase 2A p70s6k p70 S6 kinase

RL renilla luciferase

RNA ribonucleic acid

R-Smads receptor-activated Smad proteins

ROS reactive oxygen species

SAPK stress-activated protein kinase

SARA Smad anchor for receptor activation xviii SBD Smad-binding domain

SBE Smad binding element sEng soluble TGFβ co-receptor endoglin siRNA small interfering RNA

Smurf Smad ubiquitination-related factor

Sp1 stimulatory protein 1

STRAP Serine-Threonine kinase Receptor-Associated Protein

TβRI TGFβ receptor type I

TβRII TGFβ receptor type II

TGFβ transforming growth factor beta

TIF1γ transcriptional intermediary factor 1 gamma

TK thymidine kinase

TLP TRAP-1 Like Protein

TPA 12-O-tetradecanoyl phorbol-13-acetate

TRAP1 TβRI-associated protein-1

TRE TPA response element

TRIP-1 TGFβ-receptor interacting protein-1

TSP-1 thrombospondin-1

UEC untransformed epithelial cells

UTR untranslated region

YAP65 Yes-Associated Protein zOFC zebrafish ovarian follicle cells xix ACKNOWLEDGEMENTS

To my family and my previous mentors, especially my elder brother, my parents and my wife, your encouragement, support, understanding, and love have given the strength to follow my dream and to come through the tough years in my life.

I wish to express my great appreciation for my current and former thesis committee members, current and former interim committee Chairs, and former committee

Chairs, including Drs. David J. Spector, Anita K. Hopper, Sarah K. Bronson, Mark

Kester, Keith C. Cheng, Hui-Ling Chiang, Jiyue Zhu, Patrick G. Quinn, Michael F.

Verderame, Kathleen M. Mulder and Maricarmen D. Planas-Silva. I am grateful to current thesis committee for their guidence, encouragement and dedication to me to support me finish my graduate study here. I am also grateful to my former thesis committee for your expertise, constructive criticism and support, which not only guided me though my study here and will steer my future as well. Special thanks I want to give to Drs. David J. Spector, Kent E. Vrana, Anita K. Hopper, Mark Kester, Sarah K.

Bronson, Keith C. Cheng, Michael F. Verderame and Mala Chinoy. Without your support and critical guidance, I would probably not make it here now, therefore I really don’t know how to put in words to thank you, and I will keep doing research the way you guided me through my study here.

To the Genetics faculty, the faculty at the Pharmacology Department, staff and graduate student of the Genetics program and of the Pharmacology Department, I thank you all for your support and encouragement that have made my graduate study here xx memorable. Special thanks I want to give to Drs. Keith C. Cheng and Robert Levenson’s lab (especially Jessica A. Croushore) to teach and help me with the zebrafish experiments.

I’d like to thank Dan Krissinger and Rob Brucklacher at our Functional Genomics

Core Facility. Their work was a big help to my research, especially their help with

Quantitative Real-Time PCR analyses.

I would also like to acknowledge former graduate student Dr. Yangrong Zhang in

Dr. Hancock’s lab at University Park for her help with gel filtration experiments to provide valuable clues to the research in this thesis. I also want to acknowledge summer undergraduate student (Weeda Nejrabi) who helped me and contributed to the research in this thesis. 1

Chapter 1

Literature Review

1.1 Introduction

Transforming growth factor β (TGFβ) is the prototype for a superfamily of highly conserved ubiquitous peptides. More than 60 distinct TGFβ ligands have been identified, including TGFβs, Activins, bone morphogenic proteins (BMPs) and growth and differentiation factor (GDF), and phylogenetic studies have identified homologous TGFβ family members in humans and all animal model organisms (Feng and Derynck, 2005;

Newfeld et al, 1999). TGFβ signaling pathways have been shown to be essential for embryonic patterning, organogenesis, and adult tissue homeostasis, since its discovery over 20 years ago (Massague et al, 2000; Yue and Mulder, 2001; Shi and Massague,

2003). Genetic alterations in the TGFβ signaling pathways, which inactivate tumor suppressor genes or activate oncogenes, have been implicated in a vast array of human cancers and other diseases as well, highlighting the importance of TGFβ family signaling throughout the organism. For these reasons, there is a growing interest in understanding and therapeutically targeting TGFβ-mediated processes (Yingling et al, 2004; Kaklamani and Pasche, 2004; Gupta et al, 2004; Ishisaki and Matsuno, 2006). Despite advances in our understanding of the mechanisms of TGFβ signaling transduction, the mechanisms of the multifunctional TGFβ signaling is not completely clear yet. Therefore, further studies are required for deeper understanding of the underlying mechanisms of its diverse 2 biological responses. With increasing numbers of laboratories studying TGFβ signaling, additional TGFβ signaling components and pathways are likely to be discovered to mediate the diverse biological responses of this polypeptide growth factor. DYNLRB1 was identified as TGFβ receptor-interacting proteins (Ding and Mulder, 2004; Tang et al,

2002). In this thesis, the regulation of the function of DYNLRB1 and DYNLRB2 by

TGFβ, and the role of DYNLRB1 and DYNLRB2 in TGFβ-mediated signaling in mammalian cells have been investigated, as well as the function of a zebrafish homologue zDYNLRB and its role in TGFβ signaling in zebrafish ovarian follicle cells.

TGFβ family members have been shown to exert a growth inhibitory effect in most cell types (epithelial, endothelial, neuronal and haematopoietic cells) from mature tissues and hematopoietic precursor cells, which involves antiproliferation and apoptosis responses in these cells, thus to maintain tissue homeostasis under normal physiological conditions (Massague et al, 2000; Yue and Mulder, 2001; Shi and Massague, 2003;

Derynck and Zhang, 2003). However, growth inhibition is not the only biological action of TGFβ family members, since they also control various other fundamental processes, such as cell growth inhibition, migration, differentiation, apoptosis, the extracellular matrix (ECM), angiogenesis, and immune response (Massague et al, 2000; Yue and

Mulder, 2001; Shi and Massague, 2003; Derynck and Zhang, 2003). Genetic evidence from gene-ablation studies of TGFβ signaling components does not indicate a growth inhibitory effect from TGFβ during early embryogenesis (Massague et al, 2000;

Massague, 2000). For example, Dickson et al, demonstrated that the primary effect of loss of TGFβ1 in vivo in TGFβ1 null mice is not increased haematopoietic or endothelial 3 cell proliferation, but defective haematopoiesis and endothelial cell differentiation, suggesting the primary role of TGFβ1 in endothelial and haematopoietic precursors is to regulate their differentiation rather than inhibit their proliferation (Dickson et al, 1995). A variety of studies also showed that many cell types can lose the ability to respond to the growth inhibitory effect of TGFβ, and the nature of the cells and their cellular context determine together the final role of TGFβ in the regulation of growth of the cells: growth inhibition, or growth proliferation (Massague, 2000). Loss of responsiveness to growth inhibition by TGFβ in tumor cells may allow such tumor cells gain an advantage by selective inactivation TGFβ’s tumor suppressor activities, while maintaining its tumor promoting activities (Massague et al, 2000; Wakefield and Roberts, 2002). Thus, tight regulation of TGFβ cytokine signal transduction is essential for its role in maintaining adult tissue homeostasis in normal conditions and regulating normal embryonic development.

1.2 TGFβ signaling pathways

There are two types of TGFβ receptors, which are single-pass transmembrane serine/threonine kinase receptors, known as type I TGFβ receptor (TβRI) and type II

TGFβ receptor (TβRII). The TβRII kinase is constitutively active. As sown in Fig. 1,

TGFβ family ligands initiate signaling by binding and bringing together a pairs of each receptor TβRI and TβRII, to form a heterotetrameric receptor complex (Massague, 2000;

Yue and Mulder, 2001; Shi and Massague, 2003; Derynck and Zhang, 2003). This then triggers phosphorylation of intracellular signaling components, initiated by the TGFβ 4

TβRII TβRI cytoplasm P

P P Smad2/3

P Smad4

Nucleus P

P Co-activator TF

Fig. 1 Model for TGFβ regulation of transcription through Smad signaling proteins. P, phosphorylation; TF, transcription factor. 5 constitutive kinase activity of TβRII, which transphosphorylates the adjacent TβRI in the heterotetrameric receptor complex, and thereafter the activated TβRI kinases are capable of phosphorylating and activating other downstream signaling components, like the receptor-activated Smad proteins (R-Smads) (Massague, 2000; Yue and Mulder, 2001;

Shi and Massague, 2003; Derynck and Zhang, 2003). These signaling components either directly exert cytoplasmic effects or translocate into the nucleus to regulate target gene transcription. The final TGFβ signaling outcome is diverse, dependent upon the tissue and cell type, the microenvironment, and presence of other growth factors (Massague,

2000; Yue and Mulder, 2001; Roberts, 2002; Shi and Massague, 2003; Derynck and

Zhang, 2003). TGFβ signaling plays a very important role in maintaining tissue homeostasis in adults as well as in embryonic development processes. Because of such important functions, TGFβ signaling is a frequent target of dysregulation during carcinogenesis, and alteration of many signaling components have been identified in various human cancers (Levy and Hill, 2006).

TGFβ signal transduction is unique in that it activates two signaling pathways from the plasma membrane to the nucleus, the one-step amplification Smad pathway and the multiple-step amplification mitogen-activated protein kinase (MARK) pathway, to regulate its cellular and tissue activities (Massague, 2000; Yue and Mulder, 2001; Shi and

Massague, 2003). There is general agreement that most TGFβ target genes and end points are regulated by the Smad pathway, and that a few of them are regulated by the

Ras-MAPK pathway (Yue and Mulder, 2001; Piek and Roberts, 2001; Roberts, 2002;

Derynck and Zhang, 2003). In certain specific cellular contexts, TGFβ may also involve 6 other signaling pathways, like protein kinase A (PKA), protein kinase C (PKC), protein phosphatase 2A (PP2A), and phosphatidylinositol-3k-kinase (PI3K) and Rho GTPases pathways. Crosstalk exists between these pathways (Yue and Mulder, 2001; Piek and

Roberts, 2001; Roberts, 2002; Derynck and Zhang, 2003).

1.2.1 The Smad pathway

In the Smad pathway, there is only one step of signal amplification between the plasma membrane, where the TGFβ signal is received, and the nucleus, where target genes’ transcription is activated or repressed. The Smad proteins belong to a family of genes now accepted as the major known downstream signaling components of TGFβ family ligands in both invertebrates and vertebrates (Shi and Massague, 2003; Derynck and Zhang, 2003), and their nomenclature as “Smad” in the vertebrates is a merger of Sma in Caenorhabditis elegans and Mad in Drosophila melanogaster (Derynck et al, 1996). They were originally identified as downstream signaling components from genetic screens for second-site mutations to enhance the phenotype of known TGFβ signaling component mutants in Drosophila melanogaster and Caenorhabditis elegans

(Savage et al, 1996; Padgett et al, 1997). To date, three types of Smads have been identified in the family of eight mammalian Smad proteins: receptor-activated

Smads (R-Smads), which includes Smad1-3, Smad5 and Smad8; common Smad

(Co-Smad), which includes only Smad4; and inhibitory Smads (I-Smads), which includes Smad6 and 7. 7 1.2.1.1 R-Smads

Broadly speaking, R-Smads are pathway specific: Smad2 and 3 transduce signals downstream of the TGFβ/Nodal/Activin ligands, while Smad1, 5 and 8 function downstream of BMP and GDF. However, exceptions have been reported, like

TGFβ signaling through Smad1 in human breast cancer cells (Liu et al, 1998), and in intestinal epithelial cells (Yue et al, 1999a and b), and through Smad1 and

5 endothelial cells (Oh et al, 2000; Goumans et al, 2002).

The Smad2 and 3 R-Smads are phosphorylated by the activated TβRI kinases in the heterotetrameric receptor complex, and then translocate into the nucleus and regulate target gene transcription through the interaction with more than 60 nuclear proteins (Feng and Derynck, 2005). Although Smad2 and 3 are over 95% homologous and each activated by TGFβ (or Activin, etc.), recent studies showed that they have distinct functions (Felici et al, 2003; Ju et al, 2006;

Kim et al, 2005; Kretschmer et al, 2003; Kurisaki et al, 2001; Levy and Hill,

2005; Liu et al, 2003; ten Dijke and Hill, 2004; Uemura et al, 2005). However, the final signaling outcome is dependent upon other signaling pathways initiated by TGFβ as well as signaling pathways initiated by other growth factors in the specific cellular context (Yue and Mulder, 2001; Roberts, 2002; Derynck and

Zhang, 2003; Feng and Derynck, 2005).

1.2.1.2 Co-Smads

Until very recently, there has been only one mammalian Co-Smad, Smad4, 8 identified. Smad4 has had a central role, since TGFβ family signaling is very much dependent on it in that it forms complexes with all R-Smads to render them transcriptionally active (Feng and Derynck, 2005). A recent new finding by Massague and colleagues demonstrated that transcriptional intermediary factor 1γ (TIF1γ) is possibly a second Co-Smad that can compete with Smad4 and mediate different transcriptional effects (He et al, 2006). They showed that TIF1γ competes with Smad4 to bind Smad2/3 in hematopoietic, epithelial and mesenchymal cells in response to TGFβ, and that TIF1γ mediates the differentiation response of hematopoietic cells by the TGFβ-

Smad pathway, while Smad4 mediates the antiproliferative TGFβ effect on hematopoietic cells (He et al, 2006). However, since both TIF1γ and Smad4 are ubiquitously expressed, it remains to be determined whether TIF1γ regulates the differentiation of cells other than hematopoietic cells, and what role TIF1γ plays in epithelial and mesenchymal cells. TIF1γ selectively binds Smad2/3 in competition with

Smad4, but it is unknown whether similar molecules exist to bind to Smad1, 5 and 8 in competition with Smad4 for the BMP and GDF pathway.

1.2.1.3 I-Smads

I-Smads, the third type of Smads, structurally distinct from the other two types of

Smads, antagonize the Smad signaling pathway as well as other TGFβ signaling pathways. Whereas Smad6 is implicated in preferential inhibition of BMP-like signaling pathway, Smad7 is important to control TGFβ-like signaling (Nakao et al, 1997; Hata et al, 1998; Fujii et al, 1999; Ishisaki et al, 1999; Hanyu et al, 2001; Miyazono, 2000). 9 Several mechanisms have been identified so far for Smad7 to negatively regulate TGFß signaling: physical blockage of R-Smad’s phosphorylation, recruiting phosphatase such as protein phosphatase 1 (PP1) to dephosphorylate TßRI, scaffold for assembling a complex of TAK1- MKK3 and p38 MAPK to activate p38 MAPK, and recruiting ubiquitin E3 ligase such as Smurf-1 and Smurf-2 for proteasome-mediated degradation of the activated receptors (Ebisawa et al, 2001; Edlund et al, 2003; Kavsak et al, 2000; Shi et al, 2004). The expression of Smad7 is induced by TGFβ, thus forming an autoinhibitory feedback loop to terminate TGFβ signaling (Brodin et al, 2000; Nakao et al, 1997). Other cytokine signaling pathways have also been shown to induce Smad7 expression, such as the Jak/Stat signaling pathway initiated by epidermal growth factor

(EGF) and interferon-γ (Ulloa et al, 1999), and the NF-κB signaling pathway initiated by tumor growth factor-α and interleukin-1β (Bitzer et al, 2000; Nagarajan et al, 2000).

Therefore, Smad7 integrates signals from multiple signaling pathways and utilizes multiple mechanisms to negatively regulate TGFβ signaling, suggesting the central role of Smad7 as a potent natural inhibitor of TGFβ responses.

1.2.2 The MAPK pathways

The MAPK pathways also transduce signals initiated by TGFβ. MAPK pathways lead to rapid phosphorylation and activation of nuclear transcription factors in response to numerous extracellular signals. Activation of these signaling pathways results in transcriptional regulation of proteins involved in diverse cellular processes including cell proliferation, differentiation, apoptosis, cytokine production, and cytoskeletal reorganization. Three distinct groups of MAPKs have been identified in mammalian 10 cells, including the extracellular signal-regulated kinases (ERK1 and ERK2, also known as p44/p42 MAPKs), the stress-activated protein (SAP) kinases known as c-Jun N- terminal kinases (JNK1, JNK2 and JNK3), and the p38 MAPK (Yue and Mulder, 2001;

Piek and Roberts, 2001; Derynck and Zhang, 2003). These pathways incorporate sequential steps of signal amplification between the plasma membrane and the nucleus, and each step allows catalytic protein phosphorylation and activation, and amplification of the received signal, so that a much larger response can be produced from a small amount of stimulus.

TGFβ has been demonstrated in various cell types to activate ERKs (Hartsough and Mulder, 1995; Frey and Mulder, 1997; Hu et al, 1999; Ravanti et al, 1999; Funaba et al, 2002), JNKs (Frey and Mulder, 1997; Hartsough and Mulder, 1997; Atfi et al, 1997;

Engel et al, 1999; Hocevar et al, 1999; Mazars et al, 2000; Brown et al, 2002; Tian et al,

2004), and p38 MAPK (Hanafusa et al, 1999; Ravanti et al, 1999; Adachi-Yamada et al,

1999; Yu et al, 2002; Tian et al, 2004). ERK activation by TGFβ may involve Ras as upstream regulator (Yue and Mulder, 2001; Wakefield and Roberts, 2002). The rapid activation of MAPK within 5-30 minutes of TGFβ treatment was demonstrated as evidence for direct activation of MAPK by the TGFβ signaling pathway (Yue and

Mulder, 2001). More evidence were obtained from experiments employing over- expression of a dominant-negative mutant of Ras, and specific MAP/ERK kinase

(MEK1) inhibitor showing that TGFβ’s ability to activate ERK1 is blocked (Hartsough et al, 1996; Yue et al, 1999a and b), and experiments in cells over-pressing a Smad-binding defective mutant of TβRI or dominant-negative Smads and in Smad4-deficient cells 11 displaying maintained ability to activate p38 and JNK MAPK by TGFβ (Engel et al,

1999; Yu et al, 2002). Recent data suggest that the balance between the Smad pathway and the MAPK pathway as well other pathways plays an important role in the final TGFβ response, and changing the balance toward the MAPK pathway may allow cells to escape growth inhibition and apoptosis and acquire invasive and metastatic features of the tumor cells (Lehmann et al, 2000; Park et al, 2000; Yan et al, 2001; Zavadil et al, 2001; Liu et al, 2004; Tian et al, 2004). However, the in vitro feature of many such studies as well the specificity of the kinase inhibitors frequently used in such studies raised questions on the specific role of activating the MAPK pathway by TGFβ (Moustakas and Heldin, 2005).

The very recent identification of a possible alternative Co-Smad other than Smad4 for

TGFβ signaling (He et al, 2006) also raised questions on many studies previously performed in Smad4-deficient cells. Therefore, the exact mechanisms for the activation of these MAPK pathways by TGFβ are poorly understood, and remain to be explored and clarified.

1.2.3 Other signaling pathways

TGFβ has been shown to activate PKC and PKA (Choi et al, 1999; Hirota et al,

2000; Sylvia et al, 2000; Yakymovych et al, 2001), Calcium-Calmodulin-dependent protein kinase II (CamKII) (Wicks et al, 2000), PP2A (Griswold-Prenner et al, 1998;

Petritsch et al, 2000), the Rho GTPase (Edlund et al, 2002; Bhowmick et al, 2001), PI3K

(Bakin et al, 2000; Runyan et al, 2004), and NF-κB (Arsura et al, 1996; Kon et al, 1999;

Bitzer et al, 2000), etc, but their effects appear to be highly cell type specific. For example, the protein phosphatase 2A (PP2A) was recently identified due to the specific 12 interaction between its regulatory subunit Bα and the activated TβRI. It has been shown that Bα is directly phosphorylated by the activated TβRI to regulate PP2A catalytic activity to dephosphorylate and inactivate cell cycle regulatory proteins such as Akt,

ERK and p70 S6 kinase (p70s6k), and has been linked to control cell cycle progression

(Griswold-Prenner et al, 1998; Petritsch et al, 2000). TGFβ has also been shown to directly activate PI3K, as indicated by phosphorylation of its downstream signaling component Akt, and such activation has been implicated in TGFβ induced epithelial-to- mesenchymal transdifferentiation (EMT) (Bakin et al, 2000; Runyan et al, 2004;

Moustakas and Heldin, 2005). However, a lot remains to be clarified on the roles of these pathways in TGFβ-mediated responses and a deeper understanding of the mechanisms of their activation by TGFβ.

1.3 Role of TGFβ signaling in cancer and development

1.3.1 Role in cancer

It is now commonly accepted that TGFβ suppress tumorigenesis in early stages of tumor development, but promote tumor growth and metastasis in later stages (Tian et al,

2004; Wakefield and Roberts, 2002). The tumor suppressor activity of the TGFβ signaling pathway derives largely from its growth inhibitory effects in most normal cell types of adult tissues and tumor cells during the early stages of carcinogenesis. TGFβ arrest cells in the G1 phase of the cell cycle. Central to such growth inhibitory effects is its ability to downregulate the expression of c-Myc, a key cell cycle regulator, and to upregulate the expression of cyclin-dependent kinase (cdk) inhibitors p15, p21 and p27, 13 also important regulators of cell cycle (Chen et al, 2001; Claassen and Hann, 2000; Datto et al, 1995; Moustakas and Kardassis, 1998; Pardali et al, 2000; Wakefield and Roberts,

2002; Warner et al, 1999). In addition to cell cycle control, TGFβ signaling is also required for apoptosis induction and maintenance of genomic stability, etc. However, the exact mechanisms of such effects are not clear.

Because of TGFβ receptors’ importance to initiate intracellular TGFβ signaling, they naturally become frequent targets during carcinogenesis. Direct evidence that suggest involvement of disruption of TGFβ signaling in cancers actually comes from studies on the TβRII gene, TGFBR2 (Yue and Mulder, 2001; Piek and Roberts, 2001;

Levy and Hill, 2006; Sjoblom et al, 2006). Due to a 10- poly-adenine repeat in

TGFBR2 gene exon3, mutations have been found in this region in 90% of colon cancers with microsatellite instability (either hereditary non-polyposis colorectal cancer or sporadic cases), and also in about 15% of colon cancers without microsatellite instability

(Yue and Mulder, 2001; Piek and Roberts, 2001; Levy and Hill, 2006). These mutations usually result in a premature stop codon and truncated protein unable to transduce TGFβ signals. There are also other missense mutations found in TβRII in various cancers, such as E526Q, D404G and T315M (Yue and Mulder, 2001; Piek and Roberts, 2001; Levy and Hill, 2006). D404G mutation has been reported as a dominant-negative mutation by inhibiting the function of wild-type TβRII through preventing its appearance on the cell plasma membrane (Tanaka et al, 2000). But mutations are not the only way to inactivate the function of the tumor suppressor TβRII. TβRII expression is frequently decreased in various advanced cancers through epigenetic silencing of the TGFBR2 promoter (Yue 14 and Mulder, 2001; Piek and Roberts, 2001; Levy and Hill, 2006). There is strong evidence that when signaling from TβRII is decreased, growth inhibitory effect of TGFβ is selectively lost, while oncogenic effects (such as invasive growth) are promoted (Chen et al, 1993; Oft et al, 1998). This is what is exactly has been demonstrated for T315M mutation of TβRII: selectively blocking TGFβ-mediated growth arrest, while enhancing the metastatic potential of tumor cells (Yue and Mulder, 2001; Piek and Roberts, 2001).

Similarly, mutation of TβRI and epigenetic silencing of its promoter have been reported in prostate, colon, gastric and ovarian cancers (Levy and Hill, 2006; Piek and Roberts,

2001).

The major TGFβ signaling pathway, the Smad pathway, is also frequently targeted during carcinogenesis. Smad4, another tumor suppressor, was originally cloned and named as DPC4 (deleted in pancreatic cancer, locus 4). It has been frequently mutated in pancreatic and colorectal cancers, making cells lose the growth inhibitory effects of TGFβ (Hahn et al, 1996; Sjoblom et al, 2006). Subsequent studies found alteration of TGFβ signaling in almost all pancreatic cancers (Goggins et al, 1998). The tumor suppressor role of Smad4 is corroborated by observation that Smad4 heterozygous mice have increased risk for gastrointestinal cancers (Friedl et al, 1999; Taketo and

Takaku, 2000). Mutations of Smad2 have been found in sporadic colorectal carcinomas and lung cancers, and the proteins of such mutations are non-functional in transducing the growth inhibitory effects of TGFβ, thus strongly implicating the role of disruption of the

TGFβ signaling in such cancers (Eppert et al, 1996; Sjoblom et al, 2006). However, only very recently, Smad3 mutation has been found in human colorectal cancers (Sjoblom et 15 al, 2006). Smad3 knockout mice harboring a deletion in exon 2 were generated and exhibited a predisposition to colorectal adenocarcinomas, but the incidence of cancer was background dependent (Zhu et al, 1998). In contrast, other groups generated different

Smad3 knockout mice with different genetic backgrounds targeting exon 8 (Yang et al,

1999) or exon 1 (Datto et al, 1999), but have not found colon cancer in their mice. Table

1 lists the phenotypes of the TGFβ signaling component knockout mouse mutants. The reason for this discrepancy is unknown. It could be due to a gain-of-function of the

Smad3 allele that causes the colon cancer, or influences on penetrance from some genetic modifiers in the different genetic backgrounds, or something else currently unknown yet.

The embryonic lethality of the homozygous knockout mice targeting Smad2 or Smad4 prevents studies on tumorigenesis in these mice. Smad7 acts to limit the duration of

TGFβ signaling. Smad7 amplification and/or over-expression have been reported in patients with colorectal, pancreatic and endometrial cancer (Boulay et al, 2001 and 2003;

Dowdy et al, 2005; Kleeff et al, 1999), and Smad7 over-expression has also been shown to result in complete loss of TGFβ’s growth inhibitory response and enhanced tumorigenicity in pancreatic cancer, but no effect on induction of PAI-I by TGFβ (Kleeff et al, 1999). But, Smad7 knockout mouse has not been reported yet.

Overproduction of TGFβ ligand is also commonly found in many cancers, like colorectal cancer, gastric carcinoma, and prostate cancer, and has even been correlated with late stage tumor progression (Levy and Hill, 2006; Yue and Mulder, 2001). This is consistent with TGFβ’s ability to induce epithelial-mesenchymal transition, migration and invasion of tumor cells, and its paracrine effects (fibrosis, angiogenesis, 16

Table 1. Knockout mouse models of TGFβ family signaling proteins

Knock out Phenotype References mouse model Dickson et al, 1995; 50% die from yolk sac defects; 50% postnatal TGFβ1 Kulkarni et al, 1993; death from inflammatory disorders Shull et al, 1992 Craniofacial defects; skeletal defects; heart defects; TGFβ2 Sanford et al, 1997 renal defects

TGF 3 Kaartinen et al, 1995; β Cleft palate; delayed lung development Proetzel et al, 1995

Embryonic lethal at midgestation, severe defects Dong et al, 1996; TβR1 in the vascular development of the yolk sac and Elvin et al, 2000; placenta, and lacked circulating red blood cells; Larsson et al, 2001; endothelial cells enhanced cell proliferation, Larsson et al, 2003 improper migratory behavior, and impaired fibronectin production in vitro.

Disrupted hematopoesis and vasculogenesis of the Oshima et al, 1996; T R1I β yolk sac conditional knockout a lethal Leveen, et al, 2002 inflammatory

Smad2 homozygous No mesoderm development; no anteroposterior axis; Perigastrulation lethality. Nomura and Li, 1998; Defective in extra embryonic ectoderm and Waldrip et al, 1998; mesoderm induction/formation. Abnormalities in Weinstein et al, 1998. anterior-posterior axis

Severe mucosal infection and immune Datto et al, 1999; dysregulation. Colon cancer. Accelerated wound Smad3 Tomic et al, 2002; healing. Osteoporosis, osteoarthritis and Yang et al, 1999; skeletal defects. Reduced litter sizes; defective Zhu et al, 1998; immune response; accelerated wound healing; Ashcroft et al, 1999; excisional ear wound enlargement; colorectal Arany et al, 2006. tumors; defective folliculogenesis

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Knock out Phenotype References mouse model Homozygotes: Perigastrulation lethality. Defective in extraembryonic ectoderm and mesoderm induction/formation. Lack of ectodermal cell Sirard et al, 1998; Smad4 proliferation Yang et al, 1998; Xu et al, 2000 Heterozygotes: nodevelopmental abnormality, but increased proliferation of gastric polyps and tumors.

liver- Pronounced accumulation of iron in Wang et al, 2005 specific liver, kidney, and pancreas. Smad4 knockout skin specific Skin hair follicle defects and squamous cell Qiao et al, 2006 Smad4 carcinoma knockout

Smad1 mutant mice die at approximately 9.5- Smad1 10.5 days postcoitum. Extraembryonic defects: Lechleider et al, 2001; no or few primordial germ cells, lack of placenta Tremblay et al, 2001 and failure to establish a definitive embryonic circulation.

Embryonic lethal; Angiogenesis defects (enlarged Chang et al, 1999; Smad5 blood vessels surrounded by decreased numbers of vascular smooth muscle cells); mesenchymal Yang et al, 1999 apoptosis, gut, heart, and craniofacial defects.

Perinatally normal, but exhibit an age-dependent Smurf1 Yamashita et al, 2005 increase of bone mass.

Homozygous was embryonic lethal before E6.5 Dab2 due to defective cell positioning and structure Yang et al, 2002 formation of the visceral

FKBP12 Severe dilated cardiomyopathy and ventricular Shou et al, 1998 septal defects, but normal skeletal muscle. 18 inflammation and immunosuppression) on stromal tissues (Roberts, 1998).

1.3.2 Role in development

During development, the multi-functional TGFβ family signaling acts to promote the differentiation of pluripotent cells into specific cell types, and plays an important role in early embryo development processes such as axis formation (the dorsal-ventral axis, the anterior-posterior axis, and the left-right axis), tissue patterning and development of the three germ layers (ectoderm, mesoderm, and endoderm), or late development stage processes like organ specification, or the complex processes of morphogenetic movements of cells during the entire development period. A combination of genetic, embryological and molecular analyses in many model organisms (like frog, zebrafish, mouse and chicken) has provided much insight into the mechanisms of genetic control of these processes (Schier and Talbot, 2005; De Robertis et al, 2000; Kimelman and Griffin,

2000). In frog and zebrafish, many TGFβ family ligands (including Activin, Nodal,

BMPs) play important roles in dorsal-ventral patterning of the embryo. BMPs are known to promote ventral mesoderm development through a dorsal-ventral morphogen gradient across the embryo, while dorsalization of the embryo is achieved through dorsalizing factors (such as TGFβ family members Activin and Nodal), as well as the action of antagonists (like Noggin, Follistatin, and Chordin) to counter ventralizing factors (such as

BMP2) (Jones et al, 1995; Fainsod et al, 1997; Rodaway et al, 1999; Toyama et al, 1995;

Schier and Talbot, 2005). TGFβ family signaling has also been shown to play important role in dorsal-ventral patterning in Drosophila (Arora et al, 1994; Ashe and Levine, 1999; 19 Decotto and Ferguson, 2001). However, the role of TGFβ family members in dorsal- ventral axis formation has not yet been established in mouse or chicken.

The major TGFβ downstream intracellular signaling components, the Smad2, 3 and 4, are expressed ubiquitously throughout development in all adult tissues and are also essential for normal embryonic development. Their targeted knockout mouse models

(Table 1) have revealed different roles for them at different stages of embryogenesis.

Smad2 and Smad4 knockout mouse studies have shown that both are essential for gastrulation, since both knockout mice show perigastrulation lethality, and defective extraembryonic ectoderm and mesoderm induction, as well as abnormal anterior- posterior formation, although Smad4 null embryos exhibit more severe phenotype (Sirard et al, 1998; Waldrip et al, 1998; Weinstein et al, 1998; Yang et al, 1998). Since Smad2 and Smad3 are nearly identical, targeted inactivation of Smad3 gene generates surprisingly distinct phenotypes (Ashcroft et al, 1999; Datto et al, 1999; Yang et al,

1999). Two Smad3 knockout mice targeting exon 8 (Yang et al, 1999) or exon 1 (Datto et al, 1999) are viable and have been reported to display severe mucosal infection and immune dysfunction, osteoporosis and other skeletal defects (Datto et al, 1999; Yang et al, 1999), suggesting an important role for Smad3 in mucosal immune response and proper development of the skeleton. But such phenotypes have not been found in the viable Smad3 null mice harboring a deletion in exon 2 (Zhu et al, 1998), which might be due to some genetic modifiers influencing penetrance in the different genetic backgrounds or something else currently unknown yet, as indicated above. So, these studies suggest that Smad2 and Smad3 may regulate distinct TGFβ responses through 20 different target genes, despite their high homology, which is consistent with the results of functional differences between these Smads reported in cell culture studies (Kim et al,

2005; Kretschmer et al, 2003; Levy and Hill, 2005; Roberts et al, 2006).

TGFβ family ligands also activate the Ras/MAPK pathway, including p38

MAPK, JNKs and ERKs. Recent studies in zebrafish and Drosophila show that the

MAPK p38 is required specifically on the dorsal side to control the cell fate in dorsal blastomeres, and activation of p38 occurs in the future dorsal region of the embryo, but not in ventralized embryos after the dorsalizing signals are inhibited, indicating p38’s important role in dorsal-ventral axis formation (Suzanne et al, 1999; Fujii et al, 2000).

MKP3, a MAP kinase phosphatase, whose earliest expression is restricted to the future dorsal region of the embryo, was also shown to be required for dorsoventral patterning in zebrafish at the onset of gastrulation (Tsang et al, 2004). But, it remains to be determined whether p38 MAPK acts in parallel to, or crosstalk with the Smad signaling pathway to specify the initiation of dorsalizing signals, and the presence of such an intrinsic signaling pathway in other organisms.

TGFβ family signaling is also essential for organogenesis of many systems, including the nervous system and skin (Hawley et al, 1995; Bellusci et al, 1996; Zhao et al, 2000; Hogan and Kolodziej, 2002). TGFβ family members promote organogenesis by regulating a variety of cellular processes such as branching morphogenesis, EMT, cell proliferation and apoptosis (Hogan and Yingling, 1998; Hogan, 1999; Hogan and

Kolodziej, 2002). Branching morphogenesis is very important for organ development such as the lung and kidney. BMP4 is highly expressed in lung buds. BMP4 over- 21 expression in the lung buds reduces overall lung size, while blocking its signaling reduces it (Bellusci et al, 1996; Hogan and Yingling, 1998; Hogan and Kolodziej, 2002), suggesting BMP4’s important role in promoting the differentiation of lung buds into mature lung cells. TGFβ3 null mice exhibit perinatal death from unique developmental defects in the lung and palate, suggesting an essential function for TGFβ3 in the normal morphogenesis of palate and lung (Kaartinen et al, 1995). TGFβ2-null mice also exhibit perinatal mortality, but with a wide range of developmental defects, including cardiac, lung, craniofacial and urogenital defects, which involve various processes such as epithelial-mesenchymal interactions, ECM production and tissue remodeling (Sanford et al, 1997). But, surprisingly, there is no overlap on phenotypes between the knockout mice targeting different TGFβ isoforms (Table 1), indicating numerous non-compensated biological roles for these different isoforms. The processing and activation of the different TGFß isoforms may play important roles in determining such different phenotypes (Annes et al, 2003).

1.3.3 Role in ovarian follicle development

The TGFβ family signaling has profound effects on wide-ranging processes in many tissues and organ systems, including the ovary (Massague, 1998; Knight and

Glister, 2003; Pangas and Matzuk, 2004). Once established, the ovary has two major functions: first, to produce mature oocytes for fertilization, and, second, to produce hormones essential for the sexual maturation and reproductive ability of the female, as well as the development of secondary sexual characteristics. An ovarian follicle undergoes a series of complex developmental processes to become a preovulatory 22 follicle. During the process of folliculogenesis, the oocytes enlarge and mature. It has been established that the somatic cells (granulosa and theca cells) of the follicle provide a proper yet dynamic microenvironment that supports and nurtures the appropriate development of the oocyte (Eppig et al, 2002; Knight and Glister, 2003; Ge, 2005).

However, it is not completely known yet whether one cell type, either the somatic cell or the germ cell, or both determine the overall rate of follicle development (Erickson and

Shimasaki, 2000; Peng et al, 2000; Eppig et al, 2002; Ge, 2005). In recent years, exciting progress has been made towards unravelling the complex intraovarian control mechanisms. Although hormonal (such as Gonadotrophin and steroid hormones) regulation of follicle growth, apoptosis, and differentiation is critical for normal ovarian follicle development, it is the locally produced autocrine and paracrine signals in the ovarian follicles (particularly the TGFβ family signals) that allow individual follicles to grow and develop (Gougeon, 1996; Robker and Richards, 1998).

The role of the TGFβ family ligands in ovarian folliculogenesis has been studied extensively in animals. It has been demonstrated that various TGFβ family members

(TGFβ, Activin, Inhibin, GDF9, etc.) and their signaling components are expressed by ovarian somatic cells and oocytes in a developmental, stage-related manner and function as important intraovarian local regulators of folliculogenesis (McNatty et al, 1999;

Montgomery et al, 2001; Tomic et al, 2002 and 2004; Xu et al, 2002; Kohli et al, 2003 and 2005; Knight and Glister, 2003; Pangas and Matzuk, 2004). For example, all three mammalian TGFβ isoforms and their receptors are expressed in the sheep ovarian follicles, and their major intracellular signaling components Smad2 and Smad3 are 23 expressed in a stage specific manner in the rat ovarian follicle development, thus allowing different effects of the TGFβ family ligands to use the same signaling pathways during the ovarian follicle development.

Unfortunately, due to the embryonic lethality or perinatal mortality of the knockout mice targeting TGFβ1, TGFβ2, TGFβ3, TβRII, Smad2 and Smad4 (Dickson et al, 1995; Oshima et al, 1996; Kaartinen et al, 1995; Sanford et al, 1997; Weinstein et al,

1998; Yang et al, 1998), little can be gained about TGFβ signaling in ovarian follicle development. However, viable knockout mice do provide some insights about the role of

TGFβ signaling in ovarian follicle development. Smad3 knockout mice targeting exon 1, exon 2 or exon 8, are viable. Smad3 null mice targeting exon 8 exhibit slowed follicle growth, atretic follicles and degenerated oocytes, and reduced fertility, indicative of impaired ovarian follicle development and abnormal ovarian function (Tomic et al, 2002 and 2004). However, Smad3 null mice targeting exon 2 are fertile (Zhu et al, 1998).

Very recently, TGFβ1 null mice are reported to be viable and show severely impaired reproductive capacity and infertility (Ingman et al, 2006), suggesting that TGFβ1 plays a critical role in normal ovarian follicle development. Originally a different line of TGFβ1 null mice on a different genetic background was prenatally lethal (Dickson et al, 1995) or died by 3-4 weeks of age (Kulkarni et al, 1993; Shull et al, 1992). However, the reason for such different phenotypes is unknown, but may reflect differences in the genetic background of the generated knockout mice, such as influences on penetrance from genetic modifiers and genes conferring compensatory effect. Thus, for the majority of the components of the pathway, it may be necessary to generate tissue-specific 24 conditional and/or inducible knockout mouse models or to generate mouse models using the RNAi techniques in order to determine the role of the TGFβ isoforms and their signaling components during ovarian follicle development.

In vitro studies show that TGFβ regulates ovarian follicle development by modulating cell proliferation and differentiation. Although TGFβ is generally considered to be an inhibitor of cell proliferation, both inhibitory and stimulatory actions have been reported on follicle cells in vitro, depending on the species, the stage of differentiation and the presence of other growth factors (Attia et al, 2000; Coskun and Lin, 1994; Juneja et al, 1996; Lerner et al, 1995; Knight and Glister, 2003; Kohli et al, 2005; Liu et al,

1999; Pangas and Matzuk, 2004; Skinner et al, 1987). For example, TGFβ has been reported to inhibit proliferation of bovine granulosa cells harvested from large antral follicles (Skinner et al, 1987) and to prevent zebrafish premature oocyte maturation

(Kohli et al, 2003 and 2005), but it has also been reported to stimulate growth of preantral follicles dissected from adult mice ovaries (Liu et al, 1999).

Therefore, more studies are warranted to fully define the precise role of TGFβ signaling in the ovarian follicle development. However, to remember, other TGFβ family members, such as Activin, BMP, and other growth factors, such as epidermal growth factor, are operational with TGFβ signals in ovarian follicle development.

1.4 Regulation of TGFβ signaling

TGFβ family signaling is delicately regulated at multiple levels, both positively and negatively. 25 1.4.1 Regulation at level of ligand production

Ligand is the start point of the cytokine pathway, and the membrane receptor is only the midpoint of the cytokine pathway. So, the function and regulation of the cytokine pathway start with the production and secretion of each cytokine. TGFβ is not an exception. Three highly homologous isoforms of TGFβ (TGFβ1, TGFβ2, and TGFβ3, collectively referred to as TGFβ) have been cloned in mammals and localized to different human (19q13, 1q41, and 14q24, respectively) (Kim et al, 1990; Piek and

Roberts, 2001; Roberts, 1998). Expression of the three isoforms is under the control of distinct promoters. TGFβ1 promoter lacks classic TATA box and contains multiple regulatory sites that can be activated by AP1 proteins including c-Jun, c-Fos, and JunD, and various oncoproteins including Ras, Src, and HTLV1 tax, as well as Smad binding elements (SBEs) (Roberts, 1998; Mulder, 2000). TGFβ1 is the most abundant isoform in most cells and tissues, and it is most often found abnormal in disease pathogenesis, including carcinogenesis, consistent with the complex regulation of its promoters

(Roberts, 1998). TGFβ1 stimulates its own production by an autocrine manner (Roberts,

1998; Mulder, 2000). It has been demonstrated in Dr. Mulder’s laboratory that rapid activation of Ras, Erks, and JNKs in proliferating TGFβ1-sensitive untransformed epithelial cells (UECs) and human colon carcinoma cells (HCCCs) is required for AP-1 complex formation at the TGFβ1 promoter and its autocrine production (Mulder, 2000).

It also shows a TGFβ-induced AP-1 complex containing JunD, Fra-2, possibly c-Jun, and

Fos B, as well as a dependence on Smad3 for its autocrine production in UECs, and an

AP-1 complex at the proximal AP-1 site with c-Fos as the major detectable component in 26 HCCCs (Mulder, 2000; Yue and Mulder, 2000; Liu et al, 2006a). However, increased

TGFβ1 levels in most malignant cells result in further increased production of TGFβ1 by malignant cells, which are no longer inhibited by TGFβ (autocrine effects), and stimulation of the still TGFβ1-responsive stromal tissues for enhanced angiogenesis and matrix deposition (paracrine effects) (Piek and Roberts, 2001; Mulder, 2000).

Distinct from TGFβ1 promoter, TGFβ2 and TGFβ3 promoters both contain TATA boxes and a common proximal CREB binding site, which suggest hormonal and developmental control (Piek and Roberts, 2001). Very recently, it is reported in Dr.

Mulder’s laboratory that JNK and p38 MAPK activation is required for TGFβ3 autocrine production in UECs, and the major components of the TGFβ3-induced AP1 complex are

CREB1 and Smad3 (Liu et al, 2006b).

1.4.2 Regulation of ligand processing and activation

It is quite unique that, unlike many peptide growth factors that act on a restricted set of target cells, TGFβ is produced by and can act on nearly every cell type. However, only a very limited fraction of the total produced TGFβ is made mature and available for signaling, and the complexity and ubiquity of TGFβ responses is both cell type- and context-dependent, and isoform-specific (Annes et al, 2003). The release of activated mature TGFβ anywhere other than the appropriate location may produce an unwanted or even deleterious response. These all indicate a critical role of the regulation of TGFβ sequestration, processing, activation and presentation in the precise temporal and spatial control of its functions. 27 TGFβ is first synthesized as large homodimeric biologically inactive precursors.

The inactive precursor is then proteolytically processed by the mammalian convertase, furin, to yield a 25-kD homodimeric the mature protein TGFβ ligand as well as a dimeric propeptide (termed the latency-associated propeptide, LAP) (Annes et al, 2003; Beck et al, 2002; Dubois et al, 1995 and 2001). While earlier reports suggested an intracellular cleavage by furin, recent genetic evidence showed essential extracellular cleavage of the

TGFβ precursor by furin convertases (Beck et al, 2002; Zacchigna et al, 2006). Unlike most propeptides that have little affinity for the mature protein, LAP forms a complex with the mature TGFβ (called latent TGFβ complex) in which it strongly binds to the mature ligand, and thus sequesters the mature TGFβ from binding to its cell surface receptors. The LAP usually also associates with latent TGF-beta binding proteins

(LTBP) via disulfide-bound to form a large latent complex which is targeted to specific locations in the extracellular matrix by the appropriate LTBP. This latent TGFβ complex must be further processed to release the mature ligand to interact with its receptor.

1.4.2.1 Regulation of ligand availability for processing

The TGFβ precursor is proteolytically cleaved by furin to release the mature protein TGFβ ligand and the LAP. It was originally reported that an intracellular cleavage of the TGFβ precursor by furin occurs within the trans-Golgi network in the constitutive secretory pathway of almost every cell type (Dubois et al, 1995 and 2001;

Piek and Roberts, 2001). However, recent studies showed that, in addition to the Golgi apparatus, furin is also located in endosomes and on the plasma membranes in a variety of cell types including intestinal and renal epithelial cells, and endothelial cells (Mayer et 28 al, 2004; Koo et al, 2006; Teuchert et al, 1999), and genetic and biochemical evidence demonstrated essential extracellular cleavage of endogenously produced TGFβ precursors by furin to produce mature ligands (Beck et al, 2002; Koo et al, 2006).

Emilin1 (elastin microfibril interface-located protein 1) is a secreted glycoprotein associated with the ECM of blood vessels (Bressan et al, 1993). Recently, a combination of genetic and biochemical evidence showed that Emilin1 binds specifically to the immature TGFβ precursor but not the free mature TGFβ or the latent TGFβ associated with LAP, thus blocking the TGFβ precursor proteolytic processing to mature TGFβ by furin in the extracellular space (Zacchigna et al, 2006). Zacchigna et al demonstrated that

Emilin1 null mice have increased levels of active TGFβ in the vasculature ultimately leading to hypertension, and this phenotype is rescued by reduction of TGFβ gene dosage

(Zacchigna et al, 2006). It was also demonstrated that Emilin1 over-expression results in accumulation of unprocessed immature TGFβ precursors, whereas the TGFβ precursor is quickly processed in Emilin1 null cells and its stability can be restored with specific furin inhibitors, thus Emilin1 phenocoping furin inhibition (Zacchigna et al, 2006). This places Emilin1 upstream of furin in the TGFβ precursor processing in the extracellular space but downstream of its production in the cells. This also provides more evidence for essential extracellular processing of the TGFβ precursor for its maturation, and is also the most upstream step discovered so far in the regulation of the TGFβ precursor processing.

But, Emilin1 null mice develop normally (Zacchigna et al, 2006), suggesting the presence of redundant genes or modifier genes, etc. It is not clear whether Emilin1 has any role on the TGFβ precursor processing in organ systems other than the vasculature and the lung, 29 and what signal regulates the inhibition of TGFβ precursor processing by Emilin1.

Therefore, future studies are needed.

1.4.2.2 Regulation of ligand processing

Strong genetic and biochemical evidence showed that the TGFβ precursor is proteolytically processed by furin convertase either intracellularly or extracellularly

(Dubois et al, 1995 and 2001; Beck et al, 2002; Koo et al, 2006). But how this processing is regulated is unknown. However, studies showed that in various tissues

TGFβ stimulated the transcription of furin, its own converting enzyme, through crosstalk between the ERK pathway and the Smad pathway (Blanchette et al, 1997 and 2001;

Dubois et al, 2001), thus leading to augmented processing of the TGFβ precursor. But, it is not clear how the transcription of furin is negatively regulated, or how such autoinduction of its own converting enzyme by TGFβ may influence TGFβ availability in normal physiological processes and pathological conditions. Thus, clearly, more studies are required to explore these issues.

1.4.2.3 Regulation of TGFβ ligand activation

To achieve extracellular activation, the mature TGFβ ligand must be dissociated and released from the latent TGFβ complex in the ECM that renders TGFβ inactive and thus finely controls the level of mature TGFβ ligands. The temporal and spatial activation of the mature TGFβ ligand is an important step in the regulation of TGFβ signaling.

Since TGFβ is concentrated in latent TGFβ complexes to the ECM, its activation involves the disassembly of the ECM to release the mature TGFβ ligand. It has been 30 reported that latent TGFβ can be activated by various molecules, including integrins, thrombospondin-1 (TSP-1), reactive oxygen species (ROS), mild acid (pH 4.5), and proteases, which are all known to perturb the ECM environment (Annes et al, 2003).

For the activation of TGFβ in normal physiological conditions, TSP-1-mediated or integrin-mediated activation of TGFβ may be especially important, whereas activation by ROS, mild acid (pH 4.5), and proteases may be important in certain disease conditions. TSP-1 is an extracellular matrix protein and is expressed in various tissues throughout development, which is also consistent with an in vivo role as a latent TGFβ activator (Murphy-Ullrich and Poczatek, 2000). Young TSP-1 null mice demonstrate strikingly similar histological abnormalities to young TGFβ1 null mice in multiple organ systems, especially with regard to lung and pancreas pathologies, which can be reverted to wild type by treatment with TSP-1 derived peptide to activate TGFβ (Crawford et al,

1998; Murphy-Ullrich and Poczatek, 2000). Consistent with the essential role of TGFβ in normal wound healing, TSP1 null mice also exhibit altered dermal wound healing, and a decrease in active TGFβ extracted from wounds of TSP-1 null animals, while addition of active TGFβ to the TSP1 null wounds restores the wild-type phenotype (Crawford et al, 1998; Murphy-Ullrich and Poczatek, 2000). These studies indicate that a major function of TSP1 in vivo is to activate latent TGFβ. Integrin αvβ6 was first identified to control TGFβ activation specifically in epithelial cells where its low expression is restricted, while wounding or inflammation may induce the expression of vβ6 (Breuss et al, 1995; Munger et al, 1999; Miller et al, 2001). Knockout mice targeting this integrin develop exaggerated inflammation and protection from pulmonary fibrosis, which is 31 consistent with TGFβ’s role as fibrosis inducer (Border and Ruoslahti, 1992) and its induction of integrin αvβ6 for subsequent activation of itself. Therefore, this positive feedback loop is interrupted in knockout mice targeting αvβ6, thus resulting in only a minor fibrotic response in these mice (Munger et al, 1999). Recently, integrin αvß8 was reported to activate latent TGFβ, but this activation requires the activity of the membrane type 1-matrix metalloprotease (MT1-MMP) (Mu et al, 2002).

MMP-2 and MMP-9, as well as plasmin, have also been shown in vitro to activate latent TGFβ (Yu and Stamenkovic, 2000), and TGFβ has also been shown to induce the expression of MMP-2 to augment its own activation (McMahon et al, 2003). But, such a role as latent TGFβ activator in vivo has been put in doubt by studies on knockout mice targeting the genes encoding these proteases, due to the absence of any phenotype, although this might also reflect redundancy of the activating enzymes. TGFβ activation by ROS in vivo after irradiation may simply be a result of damage of the ECM and the release of mature TGFβ from the ECM during such processes. Clearly more studies are warranted to further elucidate the mechanisms of latent TGFβ activation and their role in normal physiology.

1.4.2.4 Regulation of the availability of active mature TGFβ ligand

It is known that there are antagonists to mature ligands of the TGFβ family members (Massague and Chen, 2000). For example, Follistatin is a soluble secreted glycoprotein that binds to Activin and BMPs, and antagonizes their signaling by inhibiting interaction with their corresponding receptors (de Winter et al, 1996; Iemura et 32 al, 1998). Both Chordin and Noggin bind specifically to BMPs, but not to Activin or

TGFβ, and antagonize BMP signaling by blocking BMP interaction with their receptors

(Piccolo et al, 1996). However, no such antagonist has been identified for mature TGFβ ligand until the very recent identification of a novel placenta-derived soluble TGFβ co- receptor endoglin (sEng) (Venkatesha et al, 2006). They showed that sEng blocking

TGFβ downstream signaling by preventing binding of TGFβ1 to its receptors in endothelial cells locally where active mature TGFβ1 is produced, while sEng does not impact circulating concentrations of TGFβ1 (Venkatesha et al, 2006). They also demonstrated that increased serum sEng in individuals with preeclampsia (a pregnancy- specific hypertensive syndrome) correlates with their disease severity and falls after delivery (Venkatesha et al, 2006). But how the tissue-specific production of sEng is regulated currently is unknown. Whether there are other mature TGFβ antagonists is also not clear. Future studies may give an answer. Moreover, it is reported that the α2- macroglobulin exhibits reversible binding to TGFβ with relatively low affinity, but it does not appear to compromise the biological activity of TGFβ (Vaughan and Vale,

1993). Since it is an abundant constituent of blood plasma, α2-macroglobulin may serve as a circulating reservoir and/or reduce their degradation and clearance rate.

1.4.3 Regulation of TGFβ receptor level and activity

1.4.3.1 Enhancing effects of TGFβ co-receptors

TGFβ receptors (TβRII and TβRI) are single-pass transmembrane serine/threonine kinase receptors. There are two other TGFβ co-receptors also involved 33 in TGFβ signaling, TβRIII and Endoglin. TGFβ receptor type III (TβRIII), also known as beta-glycan, is non-signaling membrane-anchored co-receptor particular for TGFβ2 isoform, since TGFβ2 itself cannot bind to TβRII to initiate the receptor complex formation and has to rely upon TβRIII to trap and present itself to TβRII to initiate the signaling. Endoglin (Eng), also known as CD105, is another cell-surface co-receptor homologous to TβRIII, and can only bind TGFβ1 and TGFβ3 (St-Jacques et al, 1994).

But, Eng cannot bind ligands by itself, and requires the presence of at least TβRII

(Barbara et al, 1999). Over-expression studies show that TβRIII enhances TGFβ responsiveness. Recently, it has been demonstrated that Eng interacts with both active and inactive TβRII, whereas it only interacts with TβRI when it is inactive, and Eng inhibits phosphorylation levels of TβRII, but increases that of TβRI, resulting in increased phosphorylation of Smad 2 but not Smad 3 and TGFβ response changes

(Guerrero-Esteo et al, 2002). Eng may be involved in the regulation of TGFβ responsiveness in vascular endothelial cells, since Eng is preferentially expressed on vascular endothelial cells (Rius et al, 1998; Botella et al, 2001). However, the mechanisms of their function remain to be elucidated.

1.4.3.2 Negatively regulating the activity of TGFβ receptors

1.4.3.2.1 Physical Blockade of TGFβ receptors

TGFβ receptors need to bind to downstream signaling components (such as

Smad2/3) to phosphorylate them, and TβRII also must bind to TβRI to transphosphorylate it. Therefore, physical blockade of such binding and/or 34 phosphorylation is naturally an important step at which to control TGFβ signaling. As mentioned above, Smad7 is a central negative regulator of TGFβ signaling in multiple ways (Ebisawa et al, 2001; Edlund et al, 2003; Kavsak et al, 2000; Shi et al, 2004). In one mechanism, Smad7 binds to the activated receptor TβRI within the TβRI/TβRII complex, and blocking the binding of R-Smads to the receptor complex, since unlike R-Smads,

Smad7 lacks the C-terminal target site for phosphorylation by TβRI and cannot be phosphorylated and released, resulting in physical blockade of TGFβ receptors activity and preventing access of R-Smads to the activated TβRI (Souchelnytskyi et al, 1998; Shi et al, 2004). Moreover, other studies identified STRAP (Serine-Threonine kinase

Receptor-Associated Protein) and YAP65 (Yes-Associated Protein) both by yeast two- hybrid systems as Smad7 partners to facilitate the recruitment of Smad7 to the activated

TβRI and stabilization of the association between Smad7 and the activated TβRI (Datta et al, 1998; Datta and Moses. 2000; Ferrigno et al, 2002; Halder et al, 2006), thus further strengthening such physical blockade of the TGFβ receptors’ activity. Such a role of

Smad7 in negatively regulating TGFβ signaling is also supported by studies on Smad7’s role on tumorigenesis and metastasis (Azuma et al, 2005; Halder et al, 2005; Kuang et al,

2006), as well as by studies on the role of its partnering proteins STRAP and YAP65 on tumorigenesis (Guo et al, 2005; Halder et al, 2006).

In addition to physically blocking intracellular signaling components' access to and phosphorylation by the TGFβ receptors, another way of physical blocking TGFβ receptor signaling is to inhibit phosphorylation and activation of TβRI by TβRII, which is also an upstream step of such negative regulation. Immunophilin FKBP12 was identified 35 using yeast two-hybrid systems and demonstrated to bind with high affinity to TβRI and to inhibit its signaling function (Wang et al, 1994). It was further demonstrated that

FKBP12 binds to the GS region of TβRI, whose phosphorylation by TβRII can activate

TβRI, and thus masks the TβRII phosphorylation sites and results in stabilization of TβRI it its inactive conformation, but does not disrupt the interaction between TβRI and TβRII

(Chen et al, 1997; Huse et al, 1999; Wang et al, 1996). Therefore TβRI cannot be activated by phosphorylation to initiate signaling. However, studies done on primary mouse embryo fibroblasts and thymocytes from FKBP12 knockout mice showed no difference in TGFβ-mediated transcriptional responses or growth inhibition from such cells from wild-type mice (Bassing et al, 1998), thus questioning the physiological relevance of the FKBP12- TβRI interaction. Therefore more studies are required to determine whether such this interaction has a physiological role in vivo.

Moreover, TRIP-1 (TGFβ-receptor interacting protein-1) has been identified to specifically interact with TβRII in a kinase-dependent way (Chen et al, 1995). TRIP-1 has also been shown to be phosphorylated on serine and threonine by the receptor kinase and to specifically repress TGFβ-mediated induction of plasminogen activator inhibitor-1

(PAI-1), without any effect on TGFβ-mediated growth inhibition, by both receptor- dependent and receptor-independent mechanisms (Choy and Derynck, 1998).

1.4.3.2.2 Enzymatically regulating the activity of TGFβ receptors

Physical blockage is only one way of Smad7 to exercise its inhibitory effects, and it is also a relatively inefficient inhibition, since by this way each Smad7 molecule can 36 only inhibit one TβRI receptor molecule. TβRI’s functions can be inhibited much more efficiently through an enzymatic approach, such as dephosphorylation of TβRI to inactivate it. The protein phosphatase 1 (PP1) is involved in TGFβ signaling, since it has been shown to be recruited to the activated TβRI and to dephosphorylate it, thus negatively regulating TGFβ signaling. Such recruitment is mediated by PP1’s regulatory/targeting subunit GADD34 (growth arrest and DNA-damage-inducible 34).

GADD34 interacts with both TßRI and Smad7 to form a TβRI–Smad7–GADD34 triple complex, which is induced by TGFβ (Hayashi et al, 1997; Nakao et al, 1997), thus terminating TGFβ signaling and limiting the duration of TGFβ signaling.

1.4.3.3 Regulating the level of TGFβ receptors

In addition to modulating the TGFβ receptor activity, the expression level of

TGFβ receptors at the cell membrane can also be regulated, which is also an efficient way of inhibiting the TGFβ signaling. However, this is irreversible way of controlling

TGFβ signaling, since it leads to terminal destruction of the receptors. Several mechanisms have been identified. The ubiquitin-mediated proteasomal degradation pathway is evolutionary conserved, and also plays an important role in control TGFβ signaling. Smurf1 and 2 (Smad ubiquitination-related factor 1 and 2) are E3 ubiquitin ligases and have been identified as Smad interacting proteins to cause the ubiquitin- mediated degradation of Smad proteins (Kavsak et al, 2000; Zhang et al, 2001; Zhu et al,

1999). Smurf1 and 2 are located in the nucleus of cells not stimulated by TGFβ, and exported into the cytoplasm together with Smad7 when cells are treated by TGFβ. 37 Through interaction with Smad7 as an adaptor protein, Smurf1 and 2 are recruited to the

TGFβ receptor complex and ubiquitinate the associated receptors, as well as Smad7, which mark them for degradation by the proteasome machinery (Ebisawa et al, 2001;

Kavsak et al, 2000; Zhang et al, 2001).

Endocytosis-lysosome pathway has long been recognized a means to terminate signaling via degradation of activated receptors after their internalization from the cell surface. TGFβ receptors have also been shown to undergo lysosomal degradation, after

TGFβ ligand binding to the TGFβ receptors and their internalization (Anders et al, 1997 and 1998; Dore et al, 1998). Recently, Di Guglielmo et al demonstrated that the TGFβ receptors are internalized through two distinct endocytic routes: the clathrin-mediated pathway, which is important for promoting signaling, and the caveolin/lipid-raft- mediated pathway, which mediates the degradation of TGFβ receptors (Di Guglielmo et al, 2003). However, receptor down-regulation can also occur via clathrin-mediated endocytosis followed by traffic to the lysosome, without caveolar involvement (Mitchell et al, 2004). Thus, differences in receptor compartmentalization and the outcome of such trafficking are likely to depend on the localization of the proteins that interact with the receptor, and may also be cell type-specific.

1.4.4 Intracellular regulation

There are a number of ways to regulate TGFβ intracellular signaling.

1.4.4.1 Regulating the Smad2/3 activity by phosphorylation

Since the major signaling pathway is the Smad pathway, the activity, level and 38 subcellular location of Smad2/3 are important targets for regulation of the TGFβ intracellular signaling. Smad2/3 are activated through phosphorylation by the activated

TβRI kinase, and translocate into the nucleus to regulate target genes’ transcription

(Massague, 2000; Feng and Derynck, 2005).

However, there are an increasing number of cellular kinases besides TβRI that can also phosphorylate Smad2/3 at distinct amino acid residues to regulate their activity.

It has been reported that cytoplasmic kinases belonging to the MAPK family can also activate Smad2/3 phosphorylation outside the C-terminal site, the TβRI phosphorylation site, (Brown et al, 1999; Engel et al, 1999). For example, activation of Smad2 via phosphorylation by active MEKK-1 (MAPK/Erk kinase kinase 1, an upstream activator of the JNK pathway) results in enhanced Smad2-Smad4 interactions, nuclear localization of Smad2 and Smad4, and selective activation of Smad2-mediated transcription in endothelial cells (Brown et al, 1999). However, the exact amino acid residue(s) phosphorylated by MEKK1 has not been identified yet. It is also reported that TGFβ- induced rapid JNK activation in a SMAD-independent manner phosphorylates Smad3 outside the TβRI phosphorylation site to facilitate both its activation by TβRI and its nuclear accumulation (Engel et al, 1999). But, the exact site of phosphorylated amino acid residues is not clear.

ERK, a member of the MAPK family, has been reported to negatively regulate the

Smad2/3 activity by phosphorylation (Kretzschmar et al, 1999). EGF, hepatocyte growth factor and oncogenic Ras can activate ERK. Activated ERK can cause phosphorylation of

Smad2/3 outside the TβRI phosphorylation site at specific residues in the region linking 39 the DNA-binding domain and the transcriptional activation domain, leading to inhibition of TGFβ-induced nuclear accumulation of Smad2/3 and Smad2/3-dependent transcription in mammary and lung epithelial cells (Kretzschmar et al, 1999). Mutation of these phosphorylation sites in Smad3 yields a Ras-resistant form that can rescue the growth inhibitory response to TGFβ in Ras-transformed cells, providing further strong evidence for a physiological role of such negative regulation by ERK.

Activation of CamKII has also been demonstrated to phosphorylate and inhibit

TGFβ-induced nuclear import and transcriptional activity of Smad2 at several serine residues (Ser110, Ser240 and Ser260), although the exact mechanism of such inhibition is not clear yet (Wicks et al, 2000). Another example is PKC, which is also demonstrated to phosphorylate several serine residues in the MH1 domain of Smad3 and thus to abrogate

DNA binding activity of Smad3 (Yakymovych et al, 2001).

1.4.4.2 Terminating the Smad2/3 signaling by dephosphorylation

Reversible phosphorylation regulates fundamental aspects of cell activity. The phosphorylation state of cellular proteins is controlled by the opposing actions of protein kinases and phosphatases. The phosphorylated proteins can be dephosphorylated by certain protein phosphatases. TGFβ-elicited phosphorylation of its major intracellular signaling components Smad2/3 is central to its cellular effects. Thus, dephosphorylation of the Smads by phosphatases is an optimal mechanism for the termination of TGFβ-

Smad signaling, although phosphorylation of Smad2/3 can be prevented or limited by physical blocking their access to the activated receptors by Smad7 and as 40 dephosphorylation of TβRI to inactivate it by phosphatases such as PP1, as mentioned above.

However, the identities of the phosphatases responsible for the R-Smads’ dephosphorylation have remained elusive until very recently (Chen et al, 2006; Lin et al,

2006). Lin et al demonstrated that PPM1A (protein phosphatase 1alpha) is a bona fide

Smad phosphatase, and dephosphorylates TGFβ-induced phosphorylated Smad2/3 (Lin et al, 2006). Both over-expression and depletion studies, as well as early embryogenesis studies in zebrafish, confirmed PPM1A’s critical role in terminating TGFβ signaling.

However, it remains to be determined whether PPM1A’s transcription and/or activity is regulated by TGFβ, or is constitutively active. It also remains to be explored whether there are more phosphatases specific for by TGFβ signaling pathway. Similarly, pyruvate dehydrogenase phosphatase has been identified by biochemical and genetic methods as a phosphatase to directly dephosphorylate a Drosophila Smad (MAD) and to dampen signal transduction of Decapentaplegic, a TGFβ family ligand in Drosophila, thus highlighting the importance of such recently identified phosphatases in TGFβ superfamily signaling (Chen et al, 2006).

1.4.4.3 Terminating the Smad2/3 signaling by irreversible degradation

In addition to the above reversible means of regulation, growing evidence suggests that Smad protein levels are also controlled by irreversible ubiquitin-mediated degradation via the proteome machinery, which may ensure a more thorough elimination of the amplified activated signals. The E3 ligases, Smurf1, Smurf2 and SCF/Roc1 have 41 been implicated in Smad degradation in unstimulated cells and stimulated cells as well

(Fukuchi et al, 2001; Lin et al, 2000; Lo and Massague, 1999).

Smurf2 has been demonstrated to be the E3 ligase responsible for activated

Smad2 degradation in the cytoplasm, and this involves the E2-conjugating enzymes

UbcH5b/c, and, to a lesser extent, Ubc3 (Lin et al, 2000; Lo and Massague, 1999).

Smurf2 exhibits higher binding affinity to activated Smad2 upon TGFβ stimulation and potently reduces the transcriptional activity of Smad2 (Lin et al, 2000; Lo and Massague,

1999). But, it has also been demonstrated that proteasomal degradation of activated

Smad2 also occur in the nucleus (Lo and Massague, 1999), however, the identity of the specific E3 ligase specifically recognizing activated Smad2 in the nucleus has not been determined yet.

Similarly, activated Smad3 has also been shown as target for ubiquitin-mediated proteasomal degradation (Fukuchi et al, 2001). The E3 ubiquitin ligase complex ROC1-

SCF has been demonstrated to induce its ubiquitination and degradation in the cytoplasm after Smad3 is exported from the nucleus to terminate Smad3 transcriptional activity

(Fukuchi et al, 2001).

1.4.4.4 Regulating the level and activity of the Co-Smad, Smad4

The common TGFβ signaling effector Smad4 has also been shown to be target for proteasomal degradation (Moren et al, 2003; Saha et al, 2001; Wan et al, 2002). Wan et al demonstrated that Jab1 antagonizes TGFβ signaling by inducing the ubiquitylation and degradation of Smad4 via the 26S proteasome (Wan et al, 2002). Saha et al showed Ras- 42 induced decrease in Smad4 expression via the MEK/ERK/MAPK pathway, which is prevented by an inhibitor of the MEK/ERK/MAPK pathway (Saha et al, 2001). Studies on Smad4 mutants derived from human cancers with missense mutation in the MH1 domain showed that all mutants exhibit enhanced polyubiquitination and proteasomal degradation (Moren et al, 2003), providing further support for the role of ubiquitination- proteasome pathway in regulating the Smad4 level in the cells. However, in all cases, the mechanism of Smad4 ubiquitination currently is not clear and remains to be determined.

In addition to regulating the level of Smad4, the regulation of Smad4 activity has been reported as well. Moren et al reported that Smad4 is mono-ubiquitinated at lysine

507 and mono- or oligo-ubiquitinated Smad4 exhibits enhanced ability to oligomerize with R-Smads and enhanced transcriptional activity (Moren et al, 2003). Recent studies show that Smad4 can also be modified by sumoylation (Lee et al, 2003; Lin et al, 2003;

Long et al, 2003 and 2004). Sumoylation, by addition of the small ubiquitin-related modifier (SUMO) to proteins, is another way to regulate protein function, like protein- protein interaction, subnuclear localization, protein-DNA interaction and enzymatic activity, but not their degradation (Schwartz and Hochstrasser, 2003). Lee et al and Lin et al reported that sumoylation of Smad4 by the SUMO E3 ligase PIASy (protein inhibitor of activated STATy) promoted the nuclear accumulation of Smad4 and enhanced Smad4 stability and Smad4-dependent transcriptional activity in mammalian cells (HeLa cells and MDA-MB-468 cells) and Xenopus embryos (Lee et al, 2003; Lin et al, 2003). However, Long et al reported the opposite effects of Smad4 SUMO modification by PIASy, a repression of Smad4 transcriptional activity in Mv1Lu cells 43 (Long et al, 2003 and 2004). Thus, it appears that the net effect of sumoylation of Smad4 may enhance or repress its transcriptional activity, depending on the target promoter analyzed and the cell type used, and perhaps the activation state of other signaling pathways.

Furthermore, TRAP1 (TβRI-associated protein-1) has been identified to interact with Smad4 in a ligand-dependent fashion and deletion constructs of TRAP1 have been shown to inhibit TGFβ signaling and diminish the interaction of Smad4 with Smad2, thus suggesting a role for TRAP1 as a specific molecular chaperone for Smad4 facilitating the

Smad2/3-Smad4 complex formation in the vicinity of the TGFβ receptors (Charng et al,

1998; Wurthner et al, 2001).

1.4.4.5 Regulating the subcellular compartmentalization of Smad2/3

Exciting new findings in a variety of cellular and developmental systems supports the proposal that intracellular trafficking and subcellular compartmentalization of signaling components can play a more direct and active role in signal propagation and amplification, and disruption of their normal trafficking to specific subcellular compartment may lead to accumulation of such signaling components at the wrong place and have devastating consequences, like cancer (Bache et al, 2004; Penheiter, 2002;

Szymkiewicz et al, 2004).

It has been demonstrated that endocytosis of TGFβ receptors and recruitment of

Smad2/3 onto EEA1-positive early endosomal compartments is required for the propagation of the TGFβ signal through the Smad proteins (Anders et al, 1997 and 1998; 44 Di Guglielmo et al, 2003; Felici et al, 2003; Hayes et al, 2002; Mitchell et al, 2004;

Runyan et al, 2005). However, the exact mechanism is not yet fully understood. Di

Guglielmo et al show that TGFβ receptors are internalized into two distinct endocytic compartments, caveolin-positive vesicles and EEA1-positive vesicles, respectively (Di

Guglielmo et al, 2003). As shown in Fig. 2, the two pathways play different roles in

TGFβ signaling through Smad2/3: the internalization into caveolin-positive vesicles promoting signaling, while the internalization into EEA1-positive vesicles dampening it

(Di Guglielmo et al, 2003).

Many adaptor proteins have been identified for Smad2/3 and have been shown to interact with Smad2/3 and to bring Smad2/3 into association with the endosomal compartments within the cell. These adaptors include SARA (Smad Anchor for Receptor

Activation) (Hu et al, 2002; Itoh et al, 2002; Tsukazaki et al, 1998), Hrs (Hepatocyte growth factor-Regulated tyrosine kinase Substrate) (Miura et al, 2000), cPML (the cytoplasmic form of the ProMyelocytic Leukemia tumor suppressor protein) (Lin et al,

2004), Dab2 (Disabled 2) (Hocevar et al, 2001 and 2005; Mishra et al, 2002), TLP

(TRAP-1 Like Protein) (Felici et al, 2003), and β-spectrin ELF (Embryonic Liver Fodrin)

(Tang et al, 2003), etc. However, so many different proteins acting as adaptor proteins for

Smad2/3 highly suggests there is functional redundancy among them, therefore, they may exhibit cell-type differences in their functions, or the presence of additional adaptor proteins for more specific interaction regulations. For example, in agreement with this, both SARA and Hrs contain a FYVE domain that binds to phosphatidylinositol 3- phosphate (PI3P) and help to bring SARA and Hrs onto EEA1-positive early endosomal 45 TGFβ Plasma Clathrin- membrane Caveolae coated pit

Caveolin-1 Smurf2 P P Smad7 P P P P P P TβRII TβRI P SARA Smad2/3

Caveolin-1 positive vesicle Early endosome

Smurf2 P P Smad7 P P P SARA P Smad2/3 P

T R degradation β TGFβ signaling

Fig. 2. TβR internalization by clathrin- and lipid-raft-mediated endocytosis. At the cell plasma membrane, the tetrameric TβR complex is internalized by two distinct endocytic pathways. The TβR complex is composed of two TβRIs and two TβRIIs. In the clathrin-mediated endocytic pathway, receptors are directly towards the early endosomes. Here, the receptors interact with SARA, which is associated with Smad2. From the early endosomes, the receptors are able to signal through Smad2 phosphorylation. In the lipid raft/caveolae-mediated endocytic pathway, TβRs associate with Smad7-Smurf2, and are internalized into caveolin-1 positive vesicles. This leads to the ubiquitin-mediated degradation of the receptors. 46 compartments, which have shown to be sites of TGFβ receptor accumulation to propagate the TGFβ signaling within the cell (Hayes et al, 2002; Hu et al, 2002; Itoh et al, 2002; Lin et al, 2004; Miura et al, 2000; Tsukazaki et al, 1998). They also both contain a Smad-binding domain (SBD) to augment TGFβ-stimulated transcriptional activity, and they seem to have a cooperative effect to enhance TGFβ signaling (Miura et al, 2000). But, Goto et al and Lu et al reported that interaction between Smad2 or 3 and

SARA is not essential for Smad2- or Smad3-dependent responses (Goto et al, 2001; Lu et al, 2002), and Miura et al also reported that the FVYE domain of Hrs is dispensable for the increased induction of TGFβ-mediated transcription. Lin et al demonstrated that cPML is required for TGFβ signaling by interacting with Smad2/3 and facilitating their recruitment onto the EEA1-positive early endosomal compartments (Lin et al, 2004).

TGFβ responses can be restored by putting back PML into the cPml-null cells or removal of the PML–RAR oncoprotein by degradation induced by retinoic acid or As2O3 treatment. However, PML is dispensable for embryogenesis, suggesting the presence of other redundant genes, or a very restricted regulatory role for cPML in the regulation of

TGFβ signaling during embryogenesis.

Growing evidence show that Smad2 and Smad3 may drive distinct TGFβ downstream signaling pathways and responses (Flanders, 2004; Ju et al, 2006; Kim et al,

2005; Kretschmer et al, 2003; Levy and Hill, 2005; Roberts et al, 2006). Consistent with these studies, Axin (Furuhashi et al, 2001), TLP (Felici et al, 2003), and β-spectrin ELF

(Tang et al, 2003) have been identified spefically as Smad3 adaptor proteins. It has been demonstrated that Axin localizes in a punctate pattern within the cell similar to that of 47 SARA, both Axin and β-spectrin ELF interact with Smad3 and their appropriate subcellular localization plays an essential role in determining their functions as Smad3 adapters facilitating its specific subcellular localization and activation by TβRI for efficient TGFβ signaling (Furuhashi et al, 2001; Tang et al, 2003). Axin mutant

(Furuhashi et al, 2001) or β-spectrin ELF deficiency (Tang et al, 2003) results in mislocalization of Smad3 and loss of the TGFβ-dependent transcriptional responses.

However, TLP interacts predominantly with TβRII, and over-expression of TLP interferes with Smad3–Smad4 complex formation and blocks the Smad3-dependent transcriptional response, while it potentiates the Smad2 response, thus suggesting a negative role of TLP in regulating the subcellular location of Smad3 (Felici et al, 2003).

However, once the TGFβ receptors are internalized, how the receptors and the signaling components (Smad2/3 and others) are directed to the appropriate subcellular localizations, another important step of regulation, is just beginning to be studied.

Previously, it has been reported that Smad2/3 are sequestered through their interaction with (MT), and TGFβ stimulation leads to their activation and release to translocate into the nucleus (Dong et al, 2000). Recently, a TGFβ receptor-interacting protein km23 was identified in Dr. Mulder’s laboratory and demonstrated to be a light chain of the motor protein dynein (Tang et al, 2002). Cytoplasmic dynein is a motor complex that transports membrane vesicles and diverse cargoes along MT in a retrograde manner (Hirokawa, 1998; King et al, 2002; Vale, 2003; Vallee et al, 2004). It plays a wide variety of functions, such as mitotic spindle assembly and orientation, positioning of the Golgi apparatus, and transport of various intracellular organelles, including 48 endosomes and lysosomes (Hirokawa, 1998; Kamal and Goldstein, 2002; King et al,

2002; Mallik and Gross, 2004; Vale, 2003; Vallee et al, 2004). The dynein motor complex is a large multimeric complex, generally consisting of heavy chains (HC), intermediate chains (IC), light-intermediate chains (LIC), light chains (LC), and other adaptor or accessory proteins, such as dynactin. Three distinct families of dynein light chains (DLCs) have been identified in mammals, including the DYNLL, DYNLT, and

DYNLRB (Pfister et al, 2005; Vale, 2003; Williams et al, 2005; Wu et al, 2005). In addition to binding the dynein intermediate chain (DIC) at distinct regions (Susalka et al,

2002), DLCs have also been shown to interact with a number of cargoes to exert diverse functions (Hirokawa, 1998; King et al, 2002; Vale, 2003; Vallee et al, 2004). km23 was cloned as a TGFβ receptor-interacting protein, and was identified also as a dynein light chain now commonly termed DYNLRB1 (Pfister et al, 2005; Tang et al, 2002).

DYNLRB1 has been shown to interact preferentially with Smad2, and is required for

Smad2-dependent signaling (Ding and Mulder, 2004; Jin et al, in revision), suggesting that TGFβ signaling components like Smad2 may be targeted by dynein motor for efficient transport along the MT for their appropriate subcellular localization, thus maximizing the efficiency of signal propagation and maintaining signal specificity.

However, the exact mechanism needs to be further investigated.

1.4.5 Transcriptional regulation of target gene expression in the nucleus

In the basal state, Smad2 and 3 are predominantly localized in the cytoplasm (Shi and Massaue, 2003). Receptor-mediated phosphorylation of Smad2 and 3 (at the C- terminal two serine residues in the SXS motif) drives their activation and induces their 49 translocation into the nucleus. Smad3 translocates into the nucleus through importinβ1 and Ran-depdendent manner (Kurisaki et al, 2001; Xiao et al, 2000), while Smad2 enters the nucleus by a different mechanism, through direct association with components of the nuclear pore complex (the nucleoporins CAN/Nup214 and Nup153) (Xu et al, 2002).

Smad4 accumulates in the nucleus by association with activated Smad2 and 3 (Kawabata et al, 1999; Liu et al, 1997; Shi and Massague, 2003). After translocation into the nucleus, in conjunction with other nuclear cofactors, Smad2 and 3 regulate the transcription of target genes.

The Smad2, 3 and 4 proteins contain two conserved structural domains, the N- terminal MH1 domain and the C-terminal MH2 domain (Shi and Massague, 2003). Both the MH1 and MH2 domains interact with a large number of proteins in the nucleus, which may modulate the function of Smad2 and 3 for transcriptional regulation. As mentioned above, Smad2 and 3 may drive distinct pathways and TGFβ responses.

Although the mechanism involved is not completely clear, recent studies have showed that it is at least partially due to their differential DNA binding capacity and unique transcriptional activation effect. As shown in Fig. 3, the major difference between

Smad2 and Smad3 is the N-terminal MH1 domain where Smad2 contains two additional stretches of amino acids whereas Smad3 does not (Yagi et al, 1999; Dennler et al, 1999).

Crystal structural studies demonstrated that Smad3 MH1 domain bound to DNA is immediately before the DNA-binding β-hairpin of the MH1 domain, while the insertion of a 30-residue stretch in the Smad2 MH1 domain disrupts the conformation of the DNA binding hairpin and precludes Smad2 from binding directly to DNA (Shi and Massague, 50

Smad3 Linker

Smad2 Exon3 Linker

MH1 domain MH2 domain

DNA binding Protein-protein interaction Smad3 MH1 binds to SBE TGFβ receptors (Smad binding element), Smad adaptors but Smad2 MH1 cannot. Transcriptional cofactors

Fig. 3. Comparison of Smad2 and Smad3 51 2003). Therefore, Smad3, but not Smad2, binds to Smad-Binding Elements (SBEs)

(termed the CAGA box), which was found in target gene promoters (including plasminogen activator inhibitor type I and Smad7 promoters) and responsible for their

TGFβ responsiveness (Dennler et al, 1998; Jonk et al, 1998). Thus, Smad3 is a transcriptional factor by itself (Dennler et al, 1998; Zawel et al, 1998), while Smad2 may function as a coactivator for other transcriptional factors like FAST-1 (Chen et al, 1996).

A broad array of transcription factors has been identified as Smad partners (Feng and Derynck, 2005). It is now generally accepted that TGFβ-mediated transcriptional regulation of target genes requires Smad2 and 3 to recruit transcriptional coactivators or corepressors to such gene promoters to elicit specific transcriptional responses

(Massague, 2000; Massague et al, 2005). Some of these factors may be ubiquitous and mediate the same response in all cell types, while other transcription factors may be cell- type specific and responsible for distinct responses (Massague et al, 2005). Through such interaction with various transcription factors, Smad proteins regulate a broad spectrum of cellular processes in a large variety of cell types, including cell proliferation, differentiation, apoptosis, and embryogenesis (Massague et al, 2000; ten Dijke et al,

2002). The MAPK pathway and other signaling pathways also transduce signals initiated by TGFβ (Moustakas and Heldin, 2005; Yue and Mulder, 2001; Derynck and Zhang,

2003), for PKC and PKA pathways (Choi et al, 1999; Hirota et al, 2000; Sylvia et al,

2000; Yakymovych et al, 2001), PI3K (Bakin et al, 2000; Runyan et al, 2004), and NF-

κB (Arsura et al, 1996; Kon et al, 1999; Bitzer et al, 2000). Although their mechanisms of activation by TGFβ are not well understood, recent studies suggest that activation of 52 these signaling pathways may phosphorylate the linker regions between the MH1 and

MH2 domains of Smad2 and 3, and allow specific crosstalk between these signaling pathways and the Smad pathway. Therefore, the Smad pathway appears to have an important function to integrate the multiple stimuli a cell receives and multiple pathways to generate specific responses.

Thus, through both positive and negative regulation at multiple levels, the multifunctional TGFβ family ligands exert diverse functions in various cellular processes, thus maintaining tissue homeostasis under normal physiological conditions. 53

Chapter 2

Requirement of DYNLRB1 for TGFβ-mediated induction of fibronectin

Dyneins are molecular motors in the cells that play a wide variety of important functions including mitotic spindle assembly and orientation, and transport of various intracellular cargoes, including organelles like endosomes. We have demonstrated previously that dynein light chain DYNLRB1 is a novel TGFβ receptor-interacting protein. Here it is demonstrated that TGFβ stimulates the phosphorylation of DYNLRB1 in Cos-1 cells and TβRII kinase is required for this phosphorylation. It is also demonstrated that this phosphorylation occurs only on serine residues of DYNLRB1, implicating DYNLRB1 as signaling intermediate downstream of TGFβ receptors. It is showed that DYNLRB1 is a cytoplasmic protein with a DYNLRB1 antibody. It has been further been demonstrated that DYNLRB1 expression knockdown using small interfering

RNA (siRNA) significantly impaired TGFβ-induced fibronectin expression in Madin

Darby canine kidney (MDCK) epithelial cells. Here, TGFβ has also been demonstrated to stimulate the recruitment of DYNLRB1 to the dynein motor complex through the DIC, which requires the TGFβ-stimulated phosphorylation of DYNLRB1, suggesting a role for

DYNLRB1 to bring specific cargo to the dynein motor. Therefore, these results indicate that DYNLRB1 is a TGFβ signaling intermediate, and is required for the induction of fibronectin expression by TGFβ. 54 2.1 INTRODUCTION

TGFβ is the prototype for a family of highly conserved ubiquitous peptides that show a remarkable diversity in the biological actions they mediate.

These biological responses include effects on cell growth, cell death, cell differentiation, and the extracellular matrix (ECM) (Derynck and Zhang, 2003;

Shi and Massague, 2003, Mehra and Wrana, 2002).

TGFβ initiates its signals by inducing a heterotetrameric receptor complex composed of TβRI and TβRII serine/threonine kinase receptors. After TGFβ binds to

TβRII, it transphosphorylates, and thereby activates TβRI. The active receptor complex then propagates signals to downstream cellular components and regulatory proteins

(Derynck and Zhang, 2003; Shi and Massague, 2003, Mehra and Wrana, 2003). Many, if not all, of the biological effects of TGF-ß are considered to be Smad-dependent, through transcriptional regulation of extracellular matrix, adhesion, and growth regulatory genes.

But, the mechanisms of these complex effects are not well understood yet. Thus, identification of additional TGFβ signaling pathways and components will assist in our understanding of the mechanisms by which alterations in these pathways contribute to human disease.

Fibronectin, a major component of the ECM, plays important roles in cell adhesion, migration, growth and differentiation (Danen and Yamada, 2001;

Pankov and Yamada, 2002). TGFβ is one of the most potent stimulators of the

ECM, and it has been shown to play a significant role in the accumulation of 55 specific ECM components such as fibronectin and collagen (Yue and Mulder,

2001; Massague et al, 2000; Roberts et al, 2003). Despite the suggestion that

Smads play a critical role in TGFβ-mediated responses, the signaling mechanisms leading to TGFβ-mediated accumulation of ECM proteins are unclear. For example, Hocevar et al have shown that TGFβ can induce fibronectin synthesis through a Jun N-terminal kinase (JNK)-dependent pathway, but Smad4 was not involved (Hocevar et al, 1999). In addition, Gooch et al reported that calcineurin was involved in TGFβ-mediated regulation of ECM accumulation (Gooch et al,

2004). It is likely that other novel TGFβ signaling intermediates are required for mediating the effects of TGFβ on the synthesis of ECM components such as fibronectin.

Dynein is a molecular motor protein that mediates intracellular transport by conveying cargo along polarized microtubules (MTs) toward the minus ends

(Hirokawa, 1998; King et al, 2002; Vale, 2003; Vallee et al, 2004). It is a large multimeric complex, generally consisting of heavy chains (HC), intermediate chains (IC), light-intermediate chains (LIC), and light chains (LC), as well as other adaptor or accessory proteins, such as dynactin. Activation of a motor may occur by posttranslational modifications, local changes in the cellular environment, or chaperone binding (Hollenbeck, 2001). Since growth factors and cytokines are known to regulate such events, the receptors and signaling pathways for these polypeptides are potential regulators of motor protein activation and 56 organelle trafficking, events which ultimately determine the collective spatial organization of the signaling pathways within the cell.

Recruitment of DLCs to the dynein complex is important not only for specifying the cargo that it binds, but also for the regulation of intracellular transport itself (Karcher et al, 2002; Vaughan and Vallee, 1995). Three distinct families of DLCs have been identified in mammals, including the DYNLL (previously termed LC8), the DYNLT

(previously termed Tctex-1/rp3), and DYNLRB (previously termed LC7/robl) (Vale,

2003; William et al, 2005; Wu et al, 2005). DYNLRB1 is a member of the DYNLRB family DLCs. We have demonstrated previously that DYNLRB1 is also a novel TGFβ receptor-interacting protein (Tang et al, 2002). In this report, it is demonstrated that

TGFβ stimulated the phosphorylation of DYNLRB1 on serine residues, and its subsequent recruitment to the dynein complex. It is also demonstrated that TβRII kinase was required for DYNLRB1 phosphorylation and interaction with DIC stimulated by

TGFβ. Further, it is demonstrated that blockade of DYNLRB1 using siRNA decreased the induction of fibronectin expression by TGFβ in MDCK cells. The results in this

Chapter suggest a role for DYNLRB1 in TGFβ signaling, and its requirement for the induction of fibronectin expression by TGFβ.

2.2 MATERIALS AND METHODS

Reagents--The anti-Flag M2 (F3165) antibody (Ab) and mouse IgG were from Sigma-

Aldrich (St. Louis, MO). The anti-dynein intermediate chain (DIC) monoclonal Ab

(MAB1618) was from Chemicon (Temecula, CA). The rabbit IgG was from Santa Cruz 57 Biotechnology, Inc. (Santa Cruz, CA). The anti-fibronectin Ab (610078) was from BD

Biosciences Transduction Laboratories (Palo Alto, CA). Protein A or G agarose were purchased from Invitrogen (Carlsbad, CA). 32P-orthophosphate (NEX-053) was from

PerkinElmer Life Sciences (Boston, MA). TGFβ1 was purchased from R & D Systems

(Minneapolis, MN). Lipofectamine 2000 (11668-019) was from Invitrogen (Carlsbad,

CA).

Antibody production--The rabbit DYNLRB1 anti-serum was prepared against the following sequence: GIPIKSTMDNPTTTQYA (corresponding to amino acids 27-43) of human DYNLRB1 (hDYNLRB1) (Strategic BioSolutions, Newark, DE, or Covance

Reseach Products, Inc, Denver, PA). Each company also provided pre-immune serum.

Cell culture--293 (CRL-1573), and COS-1 (CRL-1650) cells were purchased from

American Type Culture Collection (Manassas, VA) and were grown in DMEM supplemented with 10% FBS. MDCK cells (CCL-34) were also obtained from ATCC and were grown in MEM-α supplemented with 10% FBS. Cells with passage 10-15 are usually used for experiments. Cultures were routinely screened for mycoplasma using

Hoechst 33258 staining (5µg/ml Hanks’ solution), diluted from stock (500µg/ml) frozen at -20°C.

Transient transfections, IP/blot, Westerns, and In vivo phosphorylation assays-- were performed essentially as described previously (Hocevar et al, 1999; Tang et al,

2002; Yue and Mulder, 2000). Cells were plated in 60 cm dishe plate 24 h, prior to

transfection, and then transiently transfected with DNA constructs indicated in the 58 corresponding figures using Lipofectamine Plus (Life Technologies) according to the

manufacturer’s instructions. The cells were then harvested with lysis buffer (20 mM Tris pH 7.5, 1% Triton X-100, 10% glycerol, 137 mM NaCl, 2 mM EDTA, 25 mM - glycerophosphate, 1 mM Na3VO4 and EDTA-free complete protease inhibitor cocktail from Roche). Cell lysates were centrifuged at 14 000 g at 4°C for 15 min, followed with protein concentration measurement by BCA assay. 1000 ug cell lysates were used for immunoprecipitation with a final volume 500 ul adjusted with lysis buffer.

Immunoprecipitation samples were rolled at 4°C for at least 2 hrs or overnight, followed with adding 40 ul protein A/G Plus agarose (Santa Cruz Biotech) to each sample and incubation for at least 1 h at 4°C. Then the samples were centrifuged to remove supernatant, and wash three times with lysis buffer, each with 400 ul. Supernatants were removed from the samples and 1X loading buffer was added to each sample. The samples were boiled for 5 min, and briefly centrifuged, followed by SDS-PAGE electrophoresis and blot with appropriate antibodies. For Western blot, 50 ug cell lysates were usually used to run SDS-PAGE electrophoresis. For in vivo phosphorylation assays, cells were washed once with medium lacking phosphate, followed by 30-min incubation in the same media for depleting intracellular phosphate. Cells were then radiolabeled by incubation in the presence of 1mCi/plate [32Pi]-orthophosphate for 3h and

TGFβ was added during the last 15 min of the labeling period. Cells were then washed with 1XPBS, and lysed on ice, followed with immunoprecipitation with an anti-Flag Ab,

SDS-PAGE electrophoresis and autoradiography.

Cellular fractionation--The NE-PER Nuclear and Cytoplasmic Extraction Reagent kit 59 (78833; Pierce, Rockford, IL) was used to fractionate Mv1Lu cells according to the manufacturer’s instructions. Cells were pelleted in a microcentrifuge tube by centrifugation at 500 g for 2-3 minutes, followed by adding 200 µl ice-cold CER I to the cell pellet and vortexing the tube vigorously on the highest setting for 15 seconds to fully resuspend the cell pellet. Then the samples were incubated on ice for 10 minutes, followed by adding 11 µl ice-cold CER II to the tube and vortexing the tube twice for 5 seconds on the highest setting separated by an incubation on ice for 1 minute. Then the samples were centrifuged for 5 minutes at maximum speed in a microcentrifuge, after which the supernatant (cytoplasmic extract) fraction of the samples were immediately transferred to clean pre-chilled tubes and kept on ice or frozen at -80°C until use. Then use 100 µl ice-cold NER to resuspend the insoluble (pellet) fraction from last step, and vortex four times on the highest setting for 15 seconds, each separated by 10 min incubation on ice. Then, the samples were centrifuged at maximum speed in a microcentrifuge for 10 minutes. The supernatant (nuclear extract) fraction of the samples were immediately transferred to clean pre-chilled tubes and kept on ice or frozen at -80°C until use.

Phosphopeptide mapping and Phosphoamino acid analysis--COS-1 cells were transfected and labeled as for in vivo phosphorylation assays. After the cell lysates were normalized for radioactivity, labeled DYNLRB1 protein was immunoprecipitated with anti-Flag Ab, separated by SDS-PAGE, transferred, and visualized by autoradiography.

The membrane containing 32P-labeled DYNLRB1 was excised, and two-dimensional phosphopeptide mapping and Phosphoamino acid analysis was performed as previously described (Boyle et al, 1991; Tang et al, 2002). 60 Cell Labeling and Pulse–Chase Analysis--Studies of DYNLRB1 protein stability in cultured 293 cells that were transfected with hDYNLRB1-Flag. At 28 h posttransfection, the cells were starved for 1 h in serum-free Dulbecco's modified Eagle's medium lacking methionine and cysteine media for 1 h at 37°C, and then metabolically labeled for 45min with 300 µCi/ml L-[35S]-methionine/cysteine (Amersham Pharmacia). The pulse-labeled cells were either immediately lysed or chased for the indicated times in regular

Dulbecco's modified Eagle's medium supplemented with 10 mM cold methionine. After two gentle washes with PBS, the cells were then lysed in lysis buffer and subjected to IP using Flag Ab. The radiolabeled DYNLRB1 proteins were fractionated by 12% NuPAGE gel (Invitrogen) and visualized by autoradiography. siRNAs –siRNA plasmids were constructed by Wei Ding as follows. The sense strand of the hairpin DYNLRB1siRNA-1 corresponded to nucleotides 230-250 of the DYNLRB1 coding region (5’-AAATTATGGTTGCACCAGATA-3’). The sense strand of hairpin

DYNLRB1 siRNA-2 corresponded to nucleotides 130-150 of the DYNLRB1 coding region (5’-AACCTCATGCACAACTTCATC-3’). DYNLRB1 siRNA-1 and DYNRB siRNA-2 plasmids were transfected using Lipofectamine 2000 reagent for the fibronectin experiments. In addition, we have made a negative control siRNA (NC siRNA) plasmid, the sequence of which does not match any genes by alignment with the NCBI database using BLAST. All of these constructs contain a human RNA polymerase III U6 promoter and can form a hairpin structure in vivo that is quite similar to chemically synthesized double-stranded siRNA. The pSIREN empty vector was purchased from BD

Clontech (Cat# 631529). 61 2.3 RESULTS

2.3.1 TGFβ stimulates DYNLRB1 phosphorylation, which occurs on serine residues

We have previously identified DYNLRB1 as a TGFβ receptor-interacting protein.

Additional alignments of DYNLRB1 with sequences in the NCBI database indicated that

DYNLRB1 is the mammalian homologue of the Drosophila protein roadblock (robl), which belongs to the LC7 family of Chlamydomonas DLCs (Bowman et al, 2000).

DYNLRB/LC7/robl DLCs belong to an ancient superfamily of proteins, known as the

MglB superfamily, with representative members in all three kingdoms of life (Tang et al,

2002, Bowman et al, 2000; Koonin et al, 2000). Table 1 lists the percent homologies, identities, and similarities of the DLCs of representative members from the DYNLRB family DLCs. In addition to binding DIC at distinct regions (Susalka et al, 2002), light chains have also been shown to directly interact with a number of proteins to exert diverse functions. For example, LC8 has been reported to be a physiologically interacting partner of p21-actvivated kinase (Pak-1) (Vadlamudi et al, 2004). Pak-1 interacts with the complex of LC8 and Bim, and phosphorylates both proteins

(Vadlamudi et al, 2004). Similarly, our previous results have shown that DYNLRB1 interacts with TGFβ receptors (Tang et al, 2002).

The serine/threonine kinase activity of the TGFβ receptors mediates phosphorylation of downstream molecules to effect TGFβ responses. Thus, if

DYNLRB1 is a component of TGFβ signaling cascade, it is conceivable that the TGFβ receptors could phosphorylate DYNLRB1 as a mechanism for activation. To determine 62

Table 2: Comparision of DYNLRB1 to some other DYNLRB family members

Homologue Species % % % Amino homology identity similarity Acids ZFIN Danio rerio 75 80 93 96

DYNLRB2 Homo sapiens 70 77 91 96 robl Drosophila 67 71 81 97 melanogaster chIL7 Chlamydomonas 59 55 74 105

B15 Spematozopsis 58 56 69 98

T24H10.6 Caenorhabditis 56 47 76 95 elegans bxd Drosophila 42 23 51 101 melanogaster

LMAJFV1 Leishmania 33 33 32 103 63 whether DYNLRB1 was phosphorylated by the TGFβ receptors, in vivo phosphorylation analyses were performed after co-expression of DYNLRB1, wild-type RI and wild-type

TβRII (lanes 2, 3), or dominant negative kinase deficient form of TβRII, KNRII (lane 4) in COS-1 cells. As shown in Fig. 4A, DYNLRB1 alone was not constitutively phosphorylated (lane 1), and over-expression of both TGFβ receptors for 48 hrs with

DYNLRB1 resulted in some level of DYNLRB1 phosphorylation (lane 2). Fig. 4A showed that TGFβ treatment for 15 min enhanced DYNLRB1 phosphorylation further

(lane 3). This phosphorylation of DYNLRB1 was completely blocked when KNRII was expressed (lane 4), thereby demonstrating that the kinase activity of RII is required for

DYNLRB1 phosphorylation.

To further confirm the phosphorylation of DYNLRB1 induced by TGFβ, phosphopeptide mapping analyses were performed on phosphorylated DYNLRB1 obtained after co-expression of DYNLRB1 and both TGFβ receptors in COS-1 cells, similar to the analyses for Fig. 4A. The analyses of (32P)-labeled DYNLRB1 by two- dimensional tryptic phosphopeptide mapping showed a complex pattern of phosphopeptides both in untreated and TGFβ-treated cells, indicating multiple phosphorylation sites (Fig. 4B). A phosphopeptide (indicated by arrow), which was not detected in untreated cells, was induced upon TGFβ, and the intensity of two other phosphopeptides (indicated by arrowhead) was markedly enhanced upon TGFβ treatment. These results indicated that TGFβ treatment stimulated the phosphorylation of one site in DYNLRB1 and enhanced the phosphorylation of two other sites in

DYNLRB1. Alternatively, the phosphorylation of the three sites could be all stimulated

65 by TGFβ, because the activity of the over-expressed TGFβ receptors might be responsible for the low phosphorylation level of the two sites in DYNLRB1 in the absence of TGFβ. Collectively, the above results suggested that DYNLRB1 is a substrate for the TGFβ receptor kinase.

After complex formation, the TGFβ receptors are known to become phosphorylated on specific serine and threonine residues (Massague, 1998). Moreover,

TGFβ receptor activation effects the phosphorylation of specific serine residues in R-

Smads, which are required for TGFβ signaling (Massague, 1998). Thus, if DYNLRB1 is a substrate for the TGFβ receptor kinase activity, phosphorylation of DYNLRB1 on serine residues might be expected. In order to examine whether this was the case, phosphoamino acid analyses were performed on phosphorylated DYNLRB1 obtained in

Fig. 4A. As shown in Fig. 5, DYNLRB1 is phosphorylated primarily on serine residues, which further suggested DYNLRB1’s function as a signaling intermediate downstream of the TGFβ receptor kinases.

2.3.2 DYNLRB1 detection by rabbit polyclonal DYNLRB1 anti-serum

In order to detect endogenous DYNLRB1 protein, a rabbit polyclonal anti-serum was developed in the lab against amino acids 27-43 of hDYNLRB1 and Western blot analyses were performed on cell lysates from MDCK cells to determine whether the

DYNLRB1 anti-serum can specifically detect endogenous DYNLRB1. As indicated in

Fig. 6A, the rabbit DYNLRB1 anti-serum specifically recognized a single band of 11

68 kDa by Western blot analysis (lane 1). No band was visible when pre-immune serum was used (lane 2). Thus, the DYNLRB1 anti-serum appeared to be specific.

In order to further demonstrate the specificity of the rabbit DYNLRB1 anti-serum,

Western blot analyses were performed after subcellular fractionation of MDCK cells transfected with hDYNLRB1-Flag. As indicated in Fig. 6B, the rabbit DYNLRB1 anti- serum could detect both endogenous and exogenous DYNLRB1 protein in the cytoplasmic fractions in both the absence and presence of TGFβ (lanes 1, 3, 5, 7, 9, 11), but not in the nuclear fraction (lanes 2, 4, 6, 8, 10, 12). Other people in the lab confirmed

DYNLRB1’s cytoplasmic localization with appropriate controls (Jin et al, 2005). These results demonstrate that DYNLRB1 is a cytoplasmic protein and the rabbit anti-serum can specifically recognize both the endogenous and exogenous DYNLRB1 protein.

2.3.3 DYNLRB1 protein has a short half-life and DYNLRB1 specific siRNAs knock down its expression in MDCK cells

Next, to determine whether DYNLRB1 was a good target for knockdown by siRNA, the half-life of the DYNLRB1 protein was measured by pulse-chase experiments after transiently transfecting 293 cells with DYNLRB1-Flag. As shown in Fig. 7, pulse- chase analyses revealed that DYNLRB1 in 293 cells has a half-life of approximately 5 h, indicating DYNLRB1 knockdown by siRNA should not be difficult.

Since I had demonstrated the specificity of our DYNLRB1 Ab and the half-life of

DYNLRB1, we could now assess the knockdown effect of DYNLRB1 siRNAs on endogenous DYNLRB1 expression. To achieve this, MDCK cells were transiently

70 transfected with DYNLRB1 siRNA-1 or DYNLRB1 siRNA-2. As shown in Fig. 8A, both DYNLRB1 siRNAs induced a marked reduction in endogenous DYNLRB1 levels

(lanes 2 and 3, top panel), compared to control siRNA (lane 1, top panel).

2.3.4 DYNLRB1 knockdown reduced fibronectin induction by TGFβ in MDCK cells

Thus, it was of interest to examine whether DYNLRB1 could mediate any of the known TGFβ signaling events. TGFβ has been shown to potently induce fibronectin expression at both the mRNA and protein levels (Ignotz and Massague, 1986; Wrana et al, 1991). In addition, Hocevar et al (Hocevar et al, 1999) have shown that the JNK pathway is required for the induction of fibronectin by TGFβ. We have previously shown that over-expression of DYNLRB1 could induce JNK activation and result in phosphorylation of the downstream target c-Jun (Tang et al, 2002), suggesting that

DYNLRB1 regulates the JNK pathway and may contribute to fibronectin induction in some manner. To determine whether this was the case, the siRNA approach was chosen to examine the effect of blocking endogenous DYNLRB1 expression on induction of extracellular matrix protein fibronectin expression.

Since it has been shown that both DYNLRB1 siRNA-1 and DYNLRB1 siRNA-2 could specifically knockdown DYNRB1 in MDCK cells, Western blot analyses were performed to examine fibronectin induction by TGFβ after transient transfection of

MDCK cells with DYNLRB1 siRNA-1 and DYNLRB1 siRNA-2. As expected, TGFβ induced marked expression of fibronectin in the control siRNA transfected cells (Fig. 8B, lanes 1 and 2, top panel). In contrast, both DYNLRB1 siRNAs led to significant

72 reduction of fibronectin expression, both in the absence and presence of TGFβ (lanes 3-6, top panel). Blockade of endogenous DYNLRB1 was confirmed by blotting with the

DYNLRB1 anti-serum (bottom panel). Thus, DYNLRB1 knockdown could block fibronectin induction by both endogenous autocrine and exogenous TGFβ. These results suggest that DYNLRB1 is required for TGFβ induction of fibronectin expression.

2.3.5 TGFβ stimulates the interaction between DYNLRB1 and dynein motor, and

TβRII is required for this interaction

The above results showing that DYNLRB1 was phosphorylated after TGFβ receptors were activated and that DYNLRB1 knockdown reduced fibronectin induction by TGFβ, suggested that DYNLRB1 might function in a TGFβ signaling pathway.

Accordingly, it was of interest to determine whether TGFβ could regulate the recruitment of DYNLRB1 to the DIC. Therefore, the interaction between DYNLRB1 and DIC was examined by IP/blot analyses in MDCK cells in the absence and presence of KNRII.

When over-expressed in cells, this receptor mutant can function in a dominant-negative fashion to block the endogenous RII (Wieser et al, 1993). As shown in Fig. 4, expression of this KNRII with wild-type TβRI did not permit DYNLRB1 phosphorylation. It was demonstrated here in Fig. 9 that TGFβ induced a rapid recruitment of DYNLRB1 to the

DIC (Fig. 9, lanes 3-5, top panel). Although a basal level of interaction of DYNLRB1 and DIC was detectable in the absence of TGFβ (lane 2, top panel), the increase in the interaction between DYNLRB1 and DIC began as early as 2 min after TGFβ treatment

(lane 3, top panel). More importantly, it was demonstrated that the TGFβ-induced

74 interaction between DYNLRB1 and DIC was significantly blocked when KNRII was expressed (lanes 7-10). No specific band was detectable in EV and IgG control lanes.

Expression of DYNLRB1 and KNRII in the relevant lanes was also confirmed (middle and lower panels). Thus, TGFβ-stimulated DYNLRB1 phosphorylation is necessary for its recruitment to the dynein motor complex through the DIC and TβRII is required for this process.

2.4 DISCUSSION

DYNLRB1 was identified through its interaction with the intracellular portions of the TGFβ receptors (Tang et al, 2002). The results in this chapter demonstrate that

DYNLRB1 may play an important role for in TGFβ signaling.

It has been demonstrated here that the kinase activity of the RII receptor is required for DYNLRB1 phosphorylation, since a dominant negative kinase-deficient

TβRII (KNRII) blocked TGFβ stimulated DYNLRB1 phosphorylation. TGFβ RI did not appear to be required for DYNLRB1 phosphorylation, although RI was present in

DYNLRB1 immunoprecipitates in affinity labeling experiments (Tang et al, 2002).

Similarly, previous studies have described TGFβ signaling molecules that are regulated specifically by the RII receptors. For example, the Daxx adaptor protein has been proposed to mediate TGFβ-induced apoptosis through its interaction with RII (Perlman et al, 2001). Another example is TRIP-1, which has been identified to specifically interact with TβRII in a kinase-dependent way, and shown to be phosphorylated on serine and 75 threonine residues and involved in repression of TGFβ induced PAI-1 expression (Chen et al, 1995; Choy and Derynck. 1998).

It has also been shown that TGFβ receptor activation results in the phosphorylation of DYNLRB1 primarily on serine residues, consistent with the kinase specificity for the TGFβ receptors. For example, the R-Smads are activated by serine phosphorylation at a C-terminal SSxS motif (Souchelnytskyi et al, 1997). While this could suggest that DYNLRB1 is a direct substrate of the

TGFβ receptor kinases, phosphorylation of DYNLRB1 could be due to another kinase associated with but downstream of the TGFβ receptors. There are consensus phosphorylation sites for protein kinase A and casein kinase II within the DYNLRB1 protein. Protein kinase A and casein kinase II have recently demonstrated to be involved in TGFβ signaling (Giannouli and Kletsas, et al,

2006; Schilling and Eder, 2003; Singh and Ramji, 2006; Yang et al, 2006; Zdunek et al, 2001). To test whether protein kinase A and/or casein kinase II are responsible for the phosphorylation of DYNLRB1 or some amino acid residue phosphorylation in DYNLRB1, in vivo phosphorylation experiments may be performed for DYNLRB1 in the presence of both of TGFβ and inhibitors for the protein kinase A and/or casein kinase II (Cheusova et al, 2006; Schilling and

Eder, 2003). If protein kinase A and/or casein kinase II are responsible for the phosphorylation of DYNLRB1 or some amino acid residue phosphorylation in

DYNLRB1, it would be expected that the phosphorylation of DYNLRB1 is blocked or decreased. If this is the case, the phosphorylated amino acid residues 76 may be identified by in vitro kinase assays using the above kinase(s) on wild-type

DYNLRB1 and its site-directed mutants targeting those potential phosphorylation sites, which can be translated in vitro or expressed and purified from bacteria.

It has also been shown herein that DYNLRB1 knockdown resulted in impairment in one of major TGFβ responses, namely induction of extracellular matrix protein fibronectin expression. Since it has been demonstrated in the lab that DYNLRN1 is required for Smad2-dependent TGFβ signaling (Jin et al, in revision), and it has also been demonstrated that TGFβ-mediated induction of matrix metalloproteinase-2 is selectively dependent on Smad2 (Piek et al, 2001), it might be expected that induction of matrix metalloproteinase-2 by TGFβ be impaired after knocking down DYNLRB1. Previously it has been shown that the induction of fibronectin by TGFβ is mediated by an activation of JNK and is Smad4-independent (Hocevar et al, 1999). In addition, it has been previously shown that stable expression of DYNLRB1 resulted in a super-activation of

JNK, as well as an increase in the phosphorylation of the downstream JNK target c-Jun, suggesting that DYNLRB1 was involved in the activation of JNK by TGFβ (Tang et al,

2002). Thus, the results that DYNLRB1 is required for the induction of fibronectin by

TGFβ is consistent with previous results in the field, suggesting that DYNLRB1 may function as a signaling intermediate for the TGFβ induction of fibronectin expression, possibly via the JNK pathway.

In summary, I have provided evidence to indicate that DYNLRB1 is an important

TGFβ signaling intermediate required for TGFβ induced expression of fibronectin. 77 Future studies will unravel the precise signaling pathways and detailed mechanisms involved in the DYNLRB1-mediated TGFβ responses examined herein. 78

Chapter 3

Requirement of DYNLRB2 in Smad3-dependent TGFβ signaling

It has demonstrated in the lab that DYNLRB1 interacts with Smad2 and is required for Smad2-depdent TGFβ signaling. Human DYNLRB2 is 77% identical to human DYNLRB1. Here in this Chapter, to test the hypothesis that DYNLRB2 is involved in Smad3-dependent TGFβ signaling, the role of endogenous DYNLRB2 on

TGFβ-mediated transcriptional activation was investigated by a siRNA approach. It was demonstrated that Smad3-dependent TGFβ induction of SBE2-Luc and PAI-1 expression in HaCaT cells, and that of Smad7-Luc in Hep3B cells, is significantly impaired, after blocking endogenous DYNLRB2 expression by siRNA. However, similar blocking

DYNLRB2 expression does not inhibit the induction of ARE-Lux by TGFβ, a Smad2- dependent signaling event, thus suggesting a requirement of DYNLRB2 in Smad3- dependent TGFβ signaling. It has been shown that DYNLRB2 is present in early endosomes with Smad3 and TβRII in IEC4-1 cells in the presence of TGFβ. In addition, a preferential interaction between DYNLRB2 and Smad3 induced by TGFβ was demonstrated by immunoprecipitation/blot analyses in 293T cells and luminescence- based mammalian interaction mapping (LUMIER) analyses in IEC4-1 cells. It was shown that TGFβ stimulates a rapid recruitment of DYNLRB2 to the dynein complex, the underlying mechanism for which was demonstrated to be the TGFβ-regulated phosphorylation of DYNLRB2. Collectively, these results indicate for the first time that

DYNLRB2 is required for Smad3-dependent TGFβ signaling. 79 3.1 INTRODUCTION

TGFβ is the prototype for TGFβ superfamily of related molecules that regulate a variety of physiologic and pathologic processes, such as embryogenesis, wound healing, fibrosis, growth control, and oncogenesis (Derynck and Zhang, 2003; Mehra and Wrana,

2003; Shi and Massague, 2003). The major signaling pathway for TGFβ is the Smad signaling pathway (Mehra and Wrana, 2003; Shi and Massague, 2003). The two TGFβ pathway-restricted Smad proteins, Smad2 and Smad3, are phosphorylated by the activated TβRI kinases, and then translocated into the nucleus where they regulate target gene transcription through the interaction with more than 60 nuclear proteins (Feng and

Derynck, 2005). They share 92% identity overall in amino acid sequence, display even greater similarity in the C-terminal MH2 domain, and have some overlapping activities

(Mehra and Wrana, 2003; Shi and Massague, 2003). However, recent studies show that they may drive distinct signaling pathways and responses (Felici et al, 2003; Kurisaki et al, 2001; Liu et al, 2003; ten Dijke and Hill, 2004; Uemura et al, 2005). For example,

Smad2 and Smad3 knockout mice display distinct phenotypes. Smad3 null mice are viable and fertile, but display severe mucosal infection and immune dysfunction, osteoporosis and other skeletal defects (Datto et al, 1999; Yang et al, 1999). In contrast,

Smad2 mutant embryos exhibit perigastrulation lethality, defective extraembryonic ectoderm and mesoderm induction, and abnormal anterior-posterior formation (Waldrip et al, 1998; Weinstein et al, 1998). However, it is not completely clear how the highly homologous Smad2 and Smad3 mediate such distinct signaling events initiated by the identical ligands. 80 We have previously identified a TGFβ receptor-interacting protein DYNLRB1

(Tang et al, 2002). DYNLRB1 belongs to the DYNLRB family DLCs that have a second member, DYNLRB2, 77% identitical to DYNLRB1 in amino acid sequence. DYNLRB1 and 2 are both ubiquitously expressed, and DYNLRB2 expression is higher than

DYNLRB1, except in the liver. Jiang et al showed that DYNLRB1 is upregulated in hepatocellular carcinoma tissues, while DYNLRB2 is downregulated (Jiang et al, 2001).

DYNLRB1 is located at human 20q12-q13.11, a chromosome region frequently amplified in ovarian cancers (Jin et al, 2004; Tanner et al, 2000). In contrast,

DYNLRB2 is located in a chromosomal region (human chromosome 16q23) that is frequently lost in many tumors including hepatocellular carcinomas (Balsara et al, 2001;

Paige et al, 2000; Yakicier et al, 2001). These results suggest that DYNLRB1 and

DYNLRB2 may play different roles in tumorigenesis.

We have demonstrated previously that DYNLRB1 interacts with Smad2, but not

Smad3, and is co-localized with Smad2 at early times after TGFβ treatment (Ding and

Mulder, 2004; Jin et al, in revision; Roberts and Derynck, 2001). Blocking DYNLRB1 expression inhibits Smad2-dependent TGFβ signaling, but not Smad3-dependent TGFβ signaling (Ding and Mulder, 2004; Jin et al, in revision). Therefore, I hypothesized that

DYNLRB2 might be involved in Smad3-dependent TGFβ signaling. Here, results in this

Chapter demonstrate a distinct requirement of DYNLRB2 in Smad3-dependent TGFβ signaling, and suggest some underlying mechanisms for the involement of DYNLRB2 in

Smad3-dependent TGFβ signaling. 81 3.2 MATERIALS AND METHODS

Reagents--The anti-Flag M2 (F3165) and anti-c-myc (M5546) antibodies (Abs) and mouse IgG were from Sigma-Aldrich (St. Louis, MO). The c-myc monoclonal Ab

(9E10) developed by Dr. J. Michael Bishop was obtained from the Developmental

Studies Hybridoma Bank (Iowa City, IA). The anti-DIC monoclonal Ab was from

Chemicon (Temecula, CA). The anti-V5 Ab (R960 25) and the Lipofectamine™ 2000 transfection reagent were obtained from Invitrogen (Carlsbad, CA). The TβRII Ab (SC-

220), rabbit IgG, and protein A/G plus agarose were from Santa Cruz Biotech (Santa

Cruz, CA). 32P-orthophosphate (NEX-053) was from Perkin Elmer (Boston, MA). The rabbit Smad3 Ab (51-1500) was from Zymed (South San Francisco, CA). TGFβ1 was purchased from R & D Systems (Minneapolis, MN). The Fugene 6 transfection reagent and the anti-HA Ab (1-583-816) were from Roche Applied Science (Indianapolis, IN).

The Dual-Luciferase Reporter Assay System (Cat. # E1960) was purchased from

Promega (Madison, MI).

Cell Culture--HaCaT cells and Mv1Lu cells (CCL-64) and Hep3B cells were obtained from ATCC (Rockville, MD) and were grown in DMEM supplemented with 10% FBS.

R1B cells and DR26 cells were kindly provided by Dr. Joan Massague (Sloan-Kettering) and maintained as for Mv1Lu cells. 293T cells were obtained from T-W. Wong (Bristol-

Myers Squibb) and were maintained as for Mv1Lu cells. The rat IEC 4-1 cell line was cultured as described previously (Yue and Mulder, 2000; Tang et al, 2002). Cells with passage 10-15 are usually used for experiments. Cultures were routinely screened for 82 mycoplasma using Hoechst staining (5µg/ml Hanks’ solution), diluted from stock

(500µg/ml) frozen at -20°C.

Construction of pCMV5- DYNLRB2-Flag plasmid-- To prepare human DYNLRB2-

Flag (hDYNLRB2-Flag, hDYNLRB2 was polymerase chain reaction (PCR)-amplified from a plasmid (IMAGE cDNA clone: 781013) containing the coding region of human

DYNLRB2 (obtained from Invitrogen), with primers containing additional suitable flanking restriction enzyme sites for BglII (5’ primer) and SalI (3’ primer), and inserted into pCMV5-Flag (Sigma) after digestion with BglII and SalI restriction enzymes. The correct DNA sequences were confirmed by sequencing in both directions.

Transient Transfections, Immunoprecipitation/blot, Westerns, and In Vivo

Phosphorylation Assays-- were performed essentially as described previously (Tang et al, 2002, Hocevar et al, 2001). Cells were plated in 60 cm di she plate 24 h prior to

transfection. Cells were then transi ently transfected with DNA constructs indicated in the corresponding figures using Lipofectamine Plus or Lipofectamine 2000 transfection reagent (Life Technologies) or FuGene 6 transfection reagent (Roche) according to the

manufacturer’s instructions. The cells were then harvested with lysis buffer (20 mM Tris pH 7.5, 1% Triton X-100, 10% glycerol, 137 mM NaCl, 2 mM EDTA, 25 mM - glycerophosphate, 1 mM Na3VO4 and EDTA-free complete protease inhibitor cocktail from Roche). Cell lysates were centrifuged at 14 000 g at 4°C for 15 min, followed with protein concentration measurement by BCA assay. 1000 ug cell lysates were used for immunoprecipitation with a final volume 500 ul adjusted with lysis buffer.

Immunoprecipitation samples were rolled at 4°C for at least 2 hrs or overnight, followed 83 with adding 40 ul protein A/G Plus agarose (Santa Cruz Biotech) to each sample and incubation for at least 1 h at 4°C. Then the samples were centrifuged to remove supernatant, and wash three times with lysis buffer, each with 400 ul. Supernatants were removed from the samples and 1X loading buffer was added to each sample. The samples were boiled for 5 min, and briefly centrifuged, followed by SDS-PAGE electrophoresis and blot with appropriate antibodies. For Western blot, 50 ug cell lysates were usually used to run SDS-PAGE electrophoresis. For in vivo phosphorylation assays, cells were washed once with medium lacking phosphate, followed by 30-min incubation in the same media for depleting intracellular phosphate. Cells were then radiolabeled by incubation in the presence of 1mCi/plate [32Pi]-orthophosphate for 3h and

TGFβ was added during the last 15 min of the labeling period. Cells were then wased with 1XPBS, and lysed on ice, followed with immunoprecipitation with an anti-Flag Ab,

SDS-PAGE electrophoresis and autoradiography.

Sucrose Flotation Gradient Analyses--IEC4-1 cells were transiently transfected with hDYNLRB2-Flag. The transfection was performed with Lipofectamine™ 2000 transfection reagent following the manufacturer’s instructions. Twenty-four hours after transfection, IEC4-1 cells were washed once with serum-free (SF) medium, incubated in

SF medium for 1 h, and then incubated in the absence and presence of TGFβ for an additional 5 min. Early endosome-containing fractions were then prepared as described previously (Lin et al, 2004). In brief, IEC4-1 cells were washed twice with ice-cold PBS, scraped in PBS and centrifuged at 1,200 g at 4 °C. After addition of 5 ml homogenization buffer (250 mM sucrose, 3 mM imidazole at pH 7.4, 0.5 mM EDTA), 84 cell pellets were put on ice for 5 min, centrifuged and resuspended in 0.5 ml homogenization buffer containing the protease inhibitor cocktail (Roche). Cells were then homogenized at 4 °C by seven passages through a 25G 5/8 needle fitted onto a 1 ml plastic syringe. Homogenates were centrifuged for 10 min at 2,000 g at 4 °C and the post-nuclear supernatants were collected and brought to 40.6% sucrose using a stock solution (62% sucrose, 3 mM imidazole at pH 7.4) with a final volume of 1.1 ml and loaded at the bottom of an SW 60 centrifugation tube. A gradient consisting of three steps was then poured (1.5 ml of 35% sucrose, 3 mM imidazole at pH 7.4; 1 ml 30% sucrose, 3 mM imidazole at pH 7.4; 0.5 ml homogenization buffer). The gradient was centrifuged at 485,000 g for 90 min using an SW60 rotor. Eight 0.5 ml fractions were collected from the top of the tube, and a portion (45µl) of each was subjected to NuPAGE

(Invitrogen) analysis, followed by western blot analysis. siRNAs-- hDYNLRB2 stealth siRNA (5’-GGACAACUCAACAACUGUUCAAUAU-

3’), hDYNLRB1 stealth siRNA (5’-ACCAGAUAAAGACUAUUUCCUGAUU-3’) and scrambled stealth siRNA (5’-AAUUCUCCGAACGUGUCACGUGAGA-3’) were designed by BLOCK-iT™ RNAi Designer (Invitrogen) and synthesized by Invitrogen.

The siRNAs were transfected using Lipofectamine 2000 following its instructions.

Luciferase Reporter Assay-- HaCaT cells were plated at 1×104 cells/cm2 in 24-well plates. Twenty-four hours after plating, the cells were transfected with the indicated amounts of either hDYNLRB2 siRNA or scrambled siRNA, together with 0.1µg ARE-

Lux and 0.1µg FAST-1 (Yeo et al, 1999) or 0.1µg SBE2-Luc (Zawel et al, 1998). For the human Smad7 promoter reporter assay, Hep3B were plated at 1x104 cells/cm2 in 24- 85 well plates. Twenty-four hours after plating, the cells were transfected with the indicated amounts of either hDYNLRB2 stealth siRNA or scrambled stealth siRNA, together with

0.1µg Smad7-Luc (von Gersdorff et al, 2000). The transfection was performed with

Lipofectamine™ 2000 transfection reagent following the manufacturer’s instructions.

Renilla luciferase under the control of the SV40 promoter (pRL-SV) was used to normalize transfection efficiencies. Twenty-four hours after transfection, the cells were washed once with SF medium and incubated in SF medium for 1 h. Then the cells were cultured in the absence and presence of TGFβ (5 ng/ml) for another 18 h. The luciferase activity was measured using the Dual-Luciferase Reporter Assay System following the manufacturer’s instructions. All assays were performed in triplicate. Data are expressed as mean±SEM.

Quantitative Real-Time PCR—HaCaT cells were transfected with siRNAs as indicated.

Twenty-four hours after transfection, the medium was washed once and replaced with SF medium for 1 h, followed by incubation of the cells in the absence and presence of TGFβ

(5 ng/ml) for an additional 18 h. Total RNA was isolated from HaCaT cells using TRIzol reagent according to its instructions (Invitrogen, Carlsbad, CA), and digested with DNase

RQI (Promega) to remove any contaminating genomic DNA. RNA (18S and 28S bands) was then visualized using the Agilent 2100 Bioanalyzer (Agilent Technologies; USA) and concentrations were measured using using the NanoDrop ND-1000

Fluorospectrometer. First strand cDNA was then produced from 1.0 mg of total RNA using random hexamer primers and the SuperScript III Reverse Transcription kit

(Invitrogen; Carlsbad, CA) by standard methods. The concentration and quality of the 86 resulting cDNA was quantified and analyzed using the NanoDrop ND-1000

Fluorospectrometer. PCR amplification was performed in 96-well plate with the Applied

Biosystems Sequence Detection System 7300 at our functional genomics core facilities.

Samples were standardized to 20 ng/ul and 40 ng of cDNA per sample was then utilized as templates for real-time PCR using a SYBR Green Master Mix (Qiagen Corp, USA), in a total volume of 25 µl. The 18S rRNA primers (Eurogentec, San Diego, CA), the hDYNLRB2 primers (QT00016793, Invitrogen), and the human PAI-1 specific primers

(forward primer: 5'-GAG ACA GGC AGC TCG GAT TC-3'; reverse primer: 5'-GGC

CTC CCA AAG TGC ATT AC-3') were designed with the computer program Primer

Express (Applied Biosystems) and produced by Invitrogen (Carlsbad, CA), were also used under identical conditions. To exclude the possibility of genomic DNA contamination, control PCR reactions with no cDNA template were also included for each gene-specific primer set. Amplification data for the genes of interest was normalized to 18S within each individual PCR reaction using the relative standard curve method. Triplicates of each PCR reaction were performed and the resultant data was averaged.

LUMIER Analyses--IEC4-1 cells were transiently co-transfected with hRL-Smad2, hRL-Smad3, pRL-TK, together with Ski-Flag or DYNLRB2-Flag as indicated. pRL-TK is renilla luciferase (RL) driven by the thymidine kinase (TK) promoter and is a negative control (Barrios-Rodiles et al, 2005). Twenty-eight hours after transfection, cells were incubated in SF medium for 1h prior to incubation for 5 min in the absence and presence of TGFβ (5 ng/ml). Cells were then lysed, followed by IP using an anti-Flag Ab, or using 87 IgG as a control. Protein interactions on anti-Flag immunoprecipitates were determined by performing a renilla luciferase enzymatic assay (Barrios-Rodiles et al, 2005).

Statistical Analyses--A Student’s t-test was used to determine the level of statistical significance of luciferase assay results and Quantitative Real-Time PCR results.

3.3 RESULTS

3.3.1 DYNLRB2 siRNAs specifically block DYNLRN2 expression.

To address whether DYNLRB2 is required in Smad3-dependent TGFβ signaling, the RNAi approach was chosen to knock down endogenous DYNLRB2. First, the knockdown effect of DYNLRB2 siRNAs was determined on exogenous DYNLRB2 expression. As shown in Fig. 10A, exogenous hDYNLRB2 was detectable in the hDYNLRB2-Flag transfected cells (lane 1, top panel). Human DYNLRB2 siRNAs knocked down hDYNLRB2-Flag expression in a dose-dependent manner (lanes 2-4, top panel), while the scrambled control siRNAs did not (lanes 5-6, top panel). Equal protein loading was confirmed by blotting with a DIC Ab (bottom panel).

In order to determine whether human DYNLRB2 siRNAs can specifically knock down endogenous hDYNLRB2 expression, quantitative real-time RT-PCR analyses were performed in the absence and presence of TGFβ after transiently transfecting HaCaT with either scrambled control siRNAs, hDYNLRB1 siRNAs, or hDYNLRB2 siRNAs as indicated (Fig. 10B). As shown in Fig. 10B, transfection with hDYNLRB2 siRNAs resulted in a marked decrease of hDYNLRB2 expression with a reduction of its mRNA

90 level by approximately 50%, both in the absence and presence of TGFβ, compared to scrambled control siRNA-transfected cells and hDYNLRB1 siRNA-transfected cells. The above results indicate that human DYNLRB2 siRNAs can specifically knock down endogenous hDYNLRB2 expression.

3.3.2 DYNLRB2 knockdown significantly inhibits TGFβ induction of Smad3- dependent transcriptional activation of SBE2-Luc and Smad7-Luc.

TGFβ activates various complex cellular responses as a result of differential transcriptional regulation and nontranscriptional effects that depend upon the cell context and physiological environment (Yue and Mulder, 2001; Roberts and Wakefield, 2003;

Attisano and Wrana, 2002). The Smad proteins have been shown to be the major intracellular mediators of TGFβ signaling and regulate target gene transcription. Smad2 and Smad3 are TGFβ pathway-specific. Since we have shown that DYNLRB1 inetracts with Smad2 but not Smad3, and is required for Smad2-dependent transcriptional activation (Ding and Mulder, 2004; Jin et al, in revision; Roberts and Derynck, 2001), it was of interest to determine whether blockade of DYNLRB2 could influence Smad3- dependent transcriptional activation.

Next the hDYNLRB2 knockdown effect was determined on Smad2-dependent and Smad3-dependent transcription regulation. For this purpose, HaCaT cells were chosen, since distinct Smad2-depdent and Smad3-dependent signaling pathways have been demonstrated in HaCaT cells (Frederick et al, 2004; Kim et al, 2005; Kretschmer et al, 2003; Levy and Hill, 2005), and HaCaT cells have also been used successfully for 91 gene expression knockdown experiments by siRNAs (Kim et al, 2005; Levy and Hill,

2005). First, SBE2-Luc luciferase reporter assays were performed in the absence and presence of TGFβ, after transiently transfecting HaCaT cells with the SBE2-Luc reporter and increasing amounts of either hDYNLRB2 siRNAs or scrambled control siRNAs.

The SBE2-Luc reporter has been used previously to demonstrate a Smad3-dependent response induced by TGFβ (Zawel et al, 1998). As shown in Fig. 10C, in the scrambled control siRNA-transfected cells, TGFβ-stimulated a 7-fold induction of the SBE2-Luc reporter, but in the hDYNLRB2 siRNA-transfected cells, TGFβ-induction of SBE2-Luc activity was significantly decreased (to levels of only 3-fold), compared to that in the scrambled control siRNA-transfected cells. Such results indicate that blocking endogenous DYNLRB2 expression impaired Smad3-dependent transcriptional activation.

Smad2 and Smad3 are highly homologous and share some overlapping activities

(Mehra and Wrana, 2003; Shi and Massague, 2003), yet they have distinct functions and are regulated differentially (Felici et al, 2003; Ju et al, 2006; Kim et al, 2005; Kretschmer et al, 2003; Kurisaki et al, 2001; Levy and Hill, 2005; Liu et al, 2003; ten Dijke and Hill,

2004; Uemura et al, 2005). To address the effect of DYNLRB2 knockdown on a Smad2- dependent transcription regulation, ARE-Lux luciferase reporter assays were performed in the absence and presence of TGFβ, after transiently transfecting HaCaT cells with the

ARE-Lux reporter and FAST1, along with increasing amounts of either hDYNLRB2 siRNAs or scrambled control siRNAs. The ARE-lux reporter has previously been used to show a Smad2-dependent induction by TGFβ or Activin (Yeo et al, 1999). As shown in

Fig. 10D, both in the scrambled control siRNA-transfected cells and in the hDYNLRB2 92

HaCaT 100000

TGFβ(-) Unit) 80000 TGFβ(+)

60000

Luciferase Luciferase

-Luc Activity -Luc 2

40000 SBE

(Relative 20000

0 scrambled control siRNA 20 pmol 100 pmol - - hDYNRB2 siRNA - - 20 pmol 100 pmol

Fig. 10C. siRNA blockade of endogenous hDYNLRB2 expression results in significant inhibition of Smad3-dependent SBE2-Luc activation in HaCaT cells. HaCaT cells were transfected with increasing amount of either hDYNLRB2 siRNA or scrambled control siRNA (20 pmol/well or 100 pmol/well in 24-well plates) and 0.1 µg SBE2-Luc. 24 h after transfection, the medium was washed once and replaced with SF medium for 1 h, followed by incubation of cells in the absence and presence of TGFβ (5 ng/ml) for 18 h. Luciferase activity was measured using the Dual Luciferase Reporter Assay System. All reporter assays were performed in triplicate. 93

HaCaT 50000 45000 TGFβ(-) Units) 4000040000 TGFβ(+) 35000

Activity 3000030000

25000 Lux Lux Luciferase Luciferase 2000020000 15000

ARE- 1000010000

5000 (Relative 0 0 1 2 3 4 scrambled control siRNA 20 pmol 100 pmol - - hDYNRB2 siRNA - - 20 pmol 100 pmol

Fig. 10D. siRNA blockade of endogenous hDYNLRB2 expression has no effect on inhibition of Smad2-dependent ARE-Lux activation in HaCaT cells. Experiments were performed as for in Fig. 10B, except that 0.1 µg ARE-lux and 0.1 µg FAST-1 were used instead of SBE2-Luc. 94 siRNA-transfected cells, TGFβ stimulated approximately a 4-fold induction of the ARE-

Lux. These results indicate that blockade of DYNLRB2 had no effect on Smad2- dependent transcriptional activation.

To further investigate the DYNLRB2 knockdown effect on TGFβ-induced transcriptional activation of Smad3-dependent target gene promoter, transcription activation of the human Smad7 promoter driven Smad7-Luc was examined in Hep3B cells. Smad binding elements (SBEs) have been identified in the human Smad7 promoter and shown to be necessary for TGFβ induction (Denissova et al, 2000; von Gersdorff et al, 2000). It has been demonstrated that Smad3 and Smad4, but not Smad2, are absolutely required for induction of the Smad7 promoter by TGFβ in human MD-MBA-

468 cells (von Gersdorff et al, 2000), human Hep3B cells (Felici et al, 2003), and mouse embryonic fibroblasts (Piek et al, 2001; von Gersdorff et al, 2000). As shown in Fig.10E, in the scrambled control siRNA-transfected cells, TGFβ stimulated a 3-fold induction of the Smad7-Luc. While, in the hDYNLRB2 siRNA-transfected cells, TGFβ-induction of the Smad7-Luc was significantly decreased (to levels of only 1.5-fold). Thus, these results indicate that DYNLRB2 knockdown decreased the induction of Smad7 promoter transcriptional activation by TGFβ, further confirming the effect of blocking endogenous

DYNLRB2 expression on Smad3-dependent transcriptional activation.

3.3.3 DYNLRB2 knockdown significantly interferes with TGFβ induced PAI-1 gene expression.

To determine whether this inhibitory effect of blockade of DYNLRB2 on Smad3- dependent transcriptional activation also occurred at the level of endogenous target gene 95

Hep3B

120000 3.0 TGFβ(-) TGFβ(+)

100000 Units)

80000 1.5

60000 Luciferase Luciferase

40000 Smad7-Luc Activity Smad7-Luc

20000 (Relative 0 scrambled control siRNA + - hDYNRB2 siRNA - +

Fig. 10E. siRNA blockade of endogenous hDYNLRB2 expression inhibits TGFβ- mediated transcriptional activation of human the Smad7 promoter via Smad3 in Hep3B cells. Experiments were performed as in Fig. 10B, except that Hep3B cells were transfected with either hDYNLRB2 siRNA or scrambled control siRNA (20 pmol/well in 24-well plate) along with Smad7-Luc (0.1 µg). The fold induction by TGFβ is indicated above the bars. 96 expression, the TGFβ induced expression of plasminogen activator inhibitor-1 (PAI-1) was examined. PAI-1 is a member of a family of proteins that inhibit plasminogen activators. Once activated, plasmin provides localized protease activity in a number of physiological processess, such as cell migration (Jackson and Reidy, 1992; Majumdar et al, 2004; Perides et al, 2006; Tarui et al, 2002). It has been demonstrated that the promoter of human PAI-1 gene contains Smad3-Smad4 DNA binding sites (Dennler et al, 1998). Smad3 has been demonstrated to be an integral component of the TGFβ- mediated induction of the endogenous PAI-1 gene in many cells, including human hepatic stellate cells (Zhao et al, 2006), human keratinocyte cells HaCaT (Frederick et al,

2004), human renal mesangial cells (Song et al, 2005a; Li et al, 2006), mouse primary dermal fibroblasts (Datto et al, 1999), young adult mouse colonocytes (Mithani et al,

2004), and rat hepatic stellate cells (Liu et al, 2006d). Such PAI-1 induction is impaired by specficially blocking Smad3 function by employing either Smad3 specific inhibitor

(Liu et al, 2006d), or dominant negative Smad3 (Frederick et al, 2004), or Smad3 null cells (Datto et al, 1999; Mithani et al, 2004). For the same resons above, HaCaT cells were used here. HaCaT cells were transiently transfected with scrambled control siRNAs, hDYNLRB1 siRNAs, or hDYNLRB2 siRNAs. hDYNLRB1 siRNAs were included here as another control, since DYNLRB1 knockdown has been shown to result in a Smad2-dependent signaling impairment (Jin et al, in revision). The cells were then treated by TGFβ for 18 hrs to induce PAI-1 expression, and the PAI-1 mRNA expression levels were determined by quantitative real-time RT-PCR analyses. As shown in Fig. 11, in both scrambled control siRNA-transfected cells and hDYNLRB1 siRNA-transfected cells, PAI-1 gene expression was induced approximately 8-fold after TGFβ stimulation. 97

HaCaT 2.5 TGFβ (-) 8 8 TGFβ (+) 2.0

1.5

1.0 3 0.5

PAI-1/18S expression ratio 0 scrambled siRNA + - - hDYNLRB1 stealth siRNA - + - hDYNLRB2 stealth siRNA - - +

Fig. 11. siRNA blockade of endogenous hDYNLRB2 inhibits TGFβ-mediated induction of endogenous PAI-1 gene expression via Smad3. HaCaT cells were transfected with scrambled control siRNA, hDYNLRB1 siRNA, or hDYNLRB2 siRNA (100 pmol/well in 6-well plates). Twenty-four hours after transfection, the medium was washed once and replaced with SF medium for 1 h, followed by incubation of the cells in the absence (open bars) and presence (black bars) of TGFβ (5ng/ml) for an additional 18 h. Real-time quantitative RT-PCR analysis of human PAI-1 mRNA expression from HaCaT cells was performed as described in “Materials and Methods.” Results depict the average and standard deviation of PAI-1 mRNA levels normalized to control 18S rRNA levels, and the fold induction by TGFβ is indicated above the bars. 98 However, in hDYNLRB2 siRNA-transfected cells, TGFβ induction of PAI-1 gene expression was significantly impaired, and the induction was only about 3-fold. Thus, these results indicate that blockade of DYNLRB2 inhibits TGFβ induction of endogenous

PAI-1 gene expression. Taken together, the above results suggest that the function of

DYNLRB2 may be specific for Smad3-dependent signaling.

3.3.4 DYNLRB2 is in early endosomes with Smad3 after TGFβ stimulation.

Next, it was of interest to explore the mechanisms underlying DYNLRB2’s role in Smad3-dependent TGFβ signaling. Since previous reports have shown that TGFβ receptors, Smad 2/3 and other signaling components are localized to early endosomes for optimal signal transduction (Hayes et al, 2002; Di Guglielmo et al, 2003), it was of interest to determine whether DYNLRB2 might be co-localized with Smad3 and TGFβ receptors to early endosomes after TGFβ treatment. To assess this, sucrose flotation gradients experiments were performed to isolate the endosomal compartments enriched for early endosome antigen-1 (EEA1) as described by Lin et al (Lin et al, 2004). The bottom panel indicates the presence of EEA1 in such early endosomes fractions (Fig. 12).

As expected, in the absence of TGFβ (Fig. 12, left panels), Smad3 (2nd panel) and the majority of TβRII (top panel) were present in fractions 6-8. However, upon TGFβ stimulation (Fig. 12, right panels), the amount of TβRII and Smad3 present in EEA1- enriched fractions was increased (fractions 4-5), consistent with previous reports (Hayes et al, 2002; Di Guglielmo et al, 2003). In terms of DYNLRB2 localization (Fig. 12, 3rd panel), in the absence of TGFβ (left panel), DYNLRB2 was not present in the early

100 endosomal fractions (fractions 4-5), and instead, it accumulated in fractions 6-8.

However, as early as 5 min after TGFβ treatment (right panel), significant amount of

DYNLRB2 was relocated to the EEA1-enriched early endosomal fractions (fractions 4-

5). The results from Fig. 12 implicate that DYNLRB2 is present in early endosomes with

Smad3 and the TβRII in the presence of TGFβ.

3.3.5 TGFβ induces a preferential interaction between DYNLRB2 and Smad3.

Differential binding of Smad2 and Smad3 to other proteins has been reported previously to regulate their signaling (Ding and Mulder, 2004; Felici et al, 2003;

Furuhashi et al, 2001; Kurisaki et al, 2001; Randall et al, 2002; Remy et al, 2004;

Selvamurugan et al, 2004; Tang et al, 2003). In order to determine whether Smad3 actually interacted DYNLRB2, IP/blot analyses was performed in the absence and presence of TGFβ, after co-expression of Smad2-Myc (lanes 1-4), Smad3-Myc (lanes 5-

9), and RI-V5, RII-HA, or EV in 293T cells, with or without DYNLRB2-Flag as indicated. As shown in Fig. 13A, there was a strong interaction observed between

DYNLRB2 and Smad3 in the presence of TGFβ stimulation (lane 6), while in the absence of the TβRs or TGFβ stimulation, there was a very low level of interaction (lanes

5 and 7). In contrast, the interaction between Smad2 and DYNLRB2 was very modest even in the presence TGFβ stimulation (lane 2). Expression of RI-V5 and RII-HA without DYNLRB2 did not result in such interaction (lanes 4 and 8), and the IgG IP control was negative (lane 9). Thus, these results show that TGFβ stimulated the interaction of DYNLRB2 preferentially with Smad3.

102 To further confirm the interaction between DYNLRB2 Smad3 in a ligand- dependent manner, LUMIER analyses (Barrios-Rodiles et al, 2005) were performed in the absence and presence of TGFβ in IEC4-1 cells, after transient co-transfection with pRL-Smad2, pRL-Smad3, or pRL-TK, together with Ski-Flag or DYNLRB2-Flag as indicated. IEC4-1 cells are a rat intestinal epithelial cell line and highly responsive to

TGFβ treatment (Yue et al, 1999a; Yue and Mulder, 2001). The principle of LUMIER is as follows: one protein of interest is fused with renilla luciferase, and another protein of interest is fused with Flag tag; after co-expression of the two proteins in the cell, the cell lysates containing the two proteins were used for immunoprecipitation with an anti-Flag

Ab, or using IgG as a control. If the two proteins interact with each other, then the protein fused with renilla luciferase will be in the immunoprecipitated complex, which will give a strong signal when performing a renilla luciferase enzymatic assay on the immunoprecipitated complex (Barrios-Rodiles et al, 2005). As shown in Fig. 13B, in the absence of TGFβ, a low basal level of interaction between DYNLRB2 and Smad3 was observed (lane 7). However, addition of TGFβ resulted in an enhanced strong interaction between DYNLRB2 and Smad3 (lane 8). In contrast, no interaction between Smad2 and

DYNLRB2 was observed in the absence of TGFβ (lane 9), and there was only a low basal level of interaction of DYNLRB2 with Smad2 even in the presence of TGFβ stimulation (lane 10). As prescribed previously, the negative control pRL-TK (lanes 1-2,

5-6) was used for comparison (Barrios-Rodiles et al, 2005). As expected, there was no interaction observed between pRL and Ski or between pRL and DYNLRB2 in the absence or presence of TGFβ. The interaction between the Ski and Smad3-Flag (lanes 3- 103

IEC 4-1 pRL-TK + + - - + + - - - - - Ski-Flag + + + + ------hDYNLRB2-Flag - - - - + + + + + + + pRL-Smad2 ------+ + - pRL-Smad3 - - + + - - + + - - + TGFβ (5 min) - + - + - + - + - + +

1000000 900000 800000

Units 700000 600000 500000

Luciferase 400000 300000 200000

Relative 100000 0 1 2 3 4 5 6 7 8 9 10 11 IP Ab: Flag IgG

Fig. 13B. hDYNLRB2 interacts preferentially with Smad3 in LUMIER analyses. IEC4-1 cells were transiently co-transfected with hRL-Smad2, hRL-Smad3, or pRL-TK (negative contol), together with hDYNLRB2-Flag or Ski-Flag (positive control) as indicated. 28 h after transfection, cells were incubated in SF medium for 1h prior to incubation for 5 min in the absence (open bar) and presence of 5 ng/ml TGFβ (black bar). Cells were lysed, followed by IP using an anti-Flag Ab, or IgG as control, and analyzed as described in “Materials and Methods.” 104 4) is used as a positive control for comparison (Barrios-Rodiles et al, 2005). The IgG control was also negative (lane 11). Thus, LUMIER analyses results further confirmed the specific preferential interaction between DYNLRB2 and Smad3 in a ligand- dependent manner.

3.3.6 TGFβ stimulates the recruitment of DYNLRB2 to the dynein motor.

As mentioned earlier, DYNLRB2 is a member of the DYNLRB family of light chains of the motor dynein in mammalian cells (Ding and Mulder, 2004; Jiang et al,

2001; Jin et al, 2005; Tang et al, 2002; Pfister et al, 2005; Pfister et al, 2006). It is thought that DLCs may be important for specifying the nature of the cargo to be transported by the motor (Kamal and Goldstein, 2002; Lo et al, 2005; Mallik and Gross,

2004; Pfister, 2005; Tai et al, 1999; Wu et al, 2005). The above demonstration of a specific interaction between DYNLRB2 and Smad3 indicate that the DYNLRB2 DLC may specifically target Smad3 to the dynein motor, thus allowing its transport to its subcellular sites of action. Therefore, it is likely that extracellular signals (such as growth factors, cytokines, etc) may be able to trigger the selection of particular DLCs to be recruited to the motor in specific cellular contexts to specify and convey the cargo, such as p53 (Lo et al, 2005). Accordingly, it was of interest to determine whether TGFβ could stimulate the recruitment of the DYNLRB2 DLC to the dynein motor through DIC. For these studies, IP/blot analyses were performed using anti-DIC Ab as the IP Ab and anti-

FLAG Ab as the blotting Ab, after co-expression of RI-V5, RII-HA, and DYNLRB2-

Flag, or EV in 293T cells in the absence and presence of TGFβ. As shown in Fig. 14A,

TGFβ induced a rapid recruitment of DYNLRB2 to DIC (lanes 3-6, top panel). Although

106 a basal level of interaction of DYNLRB2 DLC and DIC was detectable (lane 3, top panel), a significant increase was visible within 5 min of TGFβ treatment (lane 5, top panel). This increase in the interaction between DYNLRB2 and DIC began as early as 5 min after TGFβ addition (lane 4, top panel) and continued to increase in a time- dependent manner (lanes 5-6, top panel). The basal level of interaction between

DYNLRB2 and DIC in the absence of TGFβ, might be due to some low level of the over- expressed TGFβ receptor activation, which is addressed in following experiments using cells with endogenous TGFβ receptors. Expression of EV only (lane 1), RI-V5 and RII-

HA (lane 2), and the IgG control (lane 7) indicated that the interaction noted was specific for DYNLRB2. Equal protein loading was confirmed by Western blotting with an anti-

DIC Ab (bottom panel). Thus, TGFβ rapidly induced the recruitment of DYNLRB2 to dynein motor through DIC.

To determine whether TGFβ could stimulate the association of DYNLRB2 with

DIC in cells expressing endogenous TGFβ receptors, IP/blot analyses were performed in

HaCaT cells. HaCaT cells express endogenous TGFβ receptors and were used in several experiments above. As shown in Fig. 14B, there was no detectable interaction between the DYNLRB2 and DIC in the absence of TGFβ (lane 2, top panel). However, TGFβ stimulation induced a time-dependent increase in the recruitment of DYNLRB2 to DIC beginning as early as 2 min after TGFβ treatment (lanes 3-7, top panel). The IgG control was negative (lane 1, top panel). Western blot analysis with anti-DIC (middle panel) and anti-Flag (bottom panel) demonstrates equal protein expression and loading. Similar results were observed in IEC 4-1 cells with endogenous TGFβ receptors (data not 107 HaCaT

hDYNRB2-Flag + + + + + + + TGFβ (min) 5 0 2 5 15 30 60 kDa 14 hDYNLRB2 6 1 2 3 4 5 6 7 IP Ab: IgG DIC Blot Ab: Flag 98 DIC 64 1 2 3 4 5 6 7 Blot Ab: DIC 14 hDYNLRB2 6 1 2 3 4 5 6 7 Blot Ab: Flag

2 1.5 units 1 0.5 Relative 0 hDYNRB2 0 2 5 15 30 60 Time on TGFβ (min)

Fig. 14B. TGFβ stimulates the recruitment of hDYNLRB2 to DIC in HaCaT cells. HaCaT cells were transiently transfected with either EV or hDYNLRB2- Flag. Thirty-two hours after transfection, cells were incubated in SF medium for 1h prior to addition of TGFβ (5 ng/ml) for the indicated times. Cells were then analyzed as in A (top panel). Western blot analysis with anti-DIC, and anti-Flag (lower panels) demonstrate equal protein loading and expression. Plot of densitometric scan of results in top panel (bottom panel). 108 shown). Collectively, these data demonstrate that TGFβ induced a rapid recruitment of

DYNLRB2 to dynein motor through DIC in three different cell lines, including two

TGFβ-responsive cell lines expressing endogenous TGFβ receptors.

3.3.7 DYNLRB2 is phosphorylated after TGFβ stimulation and TβRII is required for this phosphorylation.

Next, we wished to determine the underlying mechanism for this TGFβ-induced rapid recruitment of DYNLRB2 to dynein motor through DIC. Protein-protein interaction usually involves posttranslational modification of the interacting proteins, such as phosphorylation. TGFβ receptors display serine/threonine kinase activity and phosphorylate a number of intracellular proteins to initiate and propagate various TGFβ signaling events and responses (Yue and Mulder, 2001; Shi and Massague, 2003). For example, it has been reported that, upon stimulation with TGFβ, β-spectrin ELF are phosphorylated to induce a conformational change to facilitate its interaction with Smad3

(Tang et al, 2003). Since the above experiments showed DYNLRB2 is required for

Smad3-dependent TGFβ signaling, it is conceivable that TGFβ receptors might phosphorylate DYNLRB2 DLC to activate its recruitment to the dynein motor. First, it was of interest to determine whether DYNLRB2 could interact with the TGFβ receptors in the presence of TGFβ. For this purpose, performed IP/blot analyses was performed in the presence of TGFβ in 293T cells after transient transfection of RI-V5, RII-HA, and

DYNLRB2-Flag, or EV. As shown in Fig. 15, an interaction was observed between

DYNLRB2 and TβRII (lane 3, top panel), but no specific band was apparent after

110 expression of only EV or DYNLRB2 alone (lanes 1 and 2, top panel). The IgG control was also negative (lane 4, top panel). Expression of RI-V5, RII-HA, and DYNLRB2-

Flag was confirmed in the relevant lanes by Western blotting (two middle and one bottom panels). Thus, DYNLRB2 did interact with TβRII in the presence of TGFβ, suggesting

DYNLRB2 might be phosphorylated as a mechanism for its activation upon TGFβ receptor activation.

To determine whether TGFβ stimulation lead to DYNLRB2 phosphorylation, in vivo phosphorylation assays were performed, after transient co-expression of either

TβRII or a dominant-negative kinase deficient TβRII (KNRII) with RI and DYNLRB2 in

293T cells. The KNRII mutant receptor functions in a dominant-negative manner to block the kinase activity of endogenous TβRII when overexpressed in cells, thus blocking downstream signaling component phosphorylation and TGFβ signaling (Brand and Schneider, 1995; Feng et al, 1995; Piek and Roberts, 2001; Tang et al, 2002). As shown in Fig. 16A, there was no detectable level of DYNLRB2 phosphorylation in cells expressing both receptors for 28h with DYNLRB2 without TGFβ treatment (lane 1, top panel). However, TGFβ treatment for 15 min induced in a significant level of

DYNLRB2 phosphorylation (lane 2, top panel). Furthermore, this phosphorylation of

DYNLRB2 was completely blocked upon expression of the KNRII (lane 3, top panel).

The IgG control (lane 4, top panel) indicated that the band noted was specific for

DYNLRB2. Equal expression of DYNLRB2 was confirmed by blotting with an anti-

Flag Ab (bottom panel). Thus, the results from Fig. 16A indicate that DYNLRB2 is

112 phosphorylated upon TGFβ stimulation, and also implicate that the kinase activity of

TβRII is required for DYNLRB2 phosphorylation.

To further demonstrate the TGFβ stimulated DYNLRB2 phosphorylation and the requirement of kinase activity of TβRII and/or TβRI for DYNLRB2 phosphorylation, similar in vivo phosphorylation assays were performed in Mv1Lu cells expressing endogenous TGFβ receptors and in two receptor mutant lines derived from Mv1Lu cells,

R1B and DR26 (Boyd and massague, 1989; Laiho et al, 1990; Laiho et al, 1991). R1B cells are an Mv1Lu derivative cell line lacking TβRI, while DR26 cells are an Mv1Lu derivative cell line lacking TβRII (Laiho et al, 1990; Laiho et al, 1991). As shown in Fig.

16B, in the absence of TGFβ treatment, DYNLRB2 was not phosphorylated in the three different cell lines (lanes 1, 3, 5), further confirming that DYNLRB2 is not constitutively phosphorylated when expressed in these cells. Upon TGFβ stimulation, significant level of DYNLRB2 phosphorylation occurred in the parental Mv1Lu cells (lane 2) and similarly in R1B cells (lane 6). However, the signal was weaker in the R1B cells than in the parental Mv1Lu cells, suggesting that the presence of TβRI might enhance the phosphorylation of DYNLRB2 through TβRII, although the kinase activity of TβRI may not be required for such phosphorylation. No detectable level of DYNLRB2 phosphorylation was observed in DR26 cells in the presence of TGFβ stimulation (lane

4). Equal expression of DYNLRB2 was confirmed by blotting with an anti-Flag Ab

(bottom panel). These results provide additional strong evidence supporting that TGFβ stimulated DYNLRB2 phosphorylation and the requirement of TβRII kinase for this process.

114 3.3.8 DYNLRB2 phosphorylation mediates its rapid recruitment to the dynein motor, which is stimulated by TGFβ.

Lastly, to determine whether the TGFβ-stimulated DYNLRB2 phosphorylation play any role in mediating the TGFβ-induced rapid recruitment of DYNLRB2 to dynein motor, the interaction between DYNLRB2 and DIC was examined in HaCaT cells in the absence and presence of KNRII, which has been shown to block the phosphorylation of

DYNLRB2 (Fig. 16A). As shown in Fig. 17, in the presence of KNRII, there was barely detectable level of interaction between DYNLRB2 and DIC in the presence of TGFβ, strongly suggesting that expression of KNRII significantly blocked the interaction between DYNLRB2 and DIC stimulated by TGFβ (lanes 6-7, top panel). In contrast, in the absence of KNRII, TGFβ stimulated a rapid interaction between DYNLRB2 and DIC, as in Fig. 14B. IgG control and EV control were both negative. Equal expression and loading of DIC and DYNLRB2 were confirmed (first middle panel and bottom panel), as was the expression of KNRII in lanes 5-6 (second middle panel). Thus, these results provide strong evidence that TGFβ-stimulated phosphorylation of DYNLRB2 lead to its rapid recruitment to the dynein motor through DIC, therefore allowing its targeting

Smad3 to the dynein motor for maximizing signaling propagation efficiency and maintaining signal specificity.

3.4 DISCUSSION

We have previously shown that the DYNLRB1 plays an important role in TGFβ signaling (Chapter 2; Jin et al, 2005; Jin et al, in revision; Tang et al, 2002). To further

116 determine the function of the DYNLRB family DLCs in TGFβ signaling, a second member of this family DLC, DYNLRB2, were studied in mammalian cells in this

Chapter. Here it was demonstrated that blockade of endogenous DYNLRB2 expression impaired TGFβ stimulated Smad3-dependent transcriptional activation and target gene induction, but not Smad2-dependent transcriptional activation. Evidence was provided to support that the underlying mechanism for DYNLRB2’s role in Smad3-specific TGFβ signaling is due to TGFβ regulated specific interaction between DYNLRB2 and Smad3.

It has also been demonstrated that TGFβ stimulated a rapid recruitment of DYNLRB2 to the dynein motor, and strong evidence has been provided that the mechanism for this recruitment is due to TGFβ-induced DYNLRB2 phosphorylation, which requires the

TβRII kinase. Collectively, these results indicate for the first time that DYNLRB2 DLC is required for Smad3-dependent TGFβ signaling.

Both Smad2-dependent and Smad3-dependent transcriptional responses are central events in TGFβ signaling (Massague et al, 2000; Moustakas et al, 2001; Shi and

Massague, 2003; ten Dijke and Hill, 2004). However, more recent evidence suggests that these two highly homologous Smad proteins play unique roles downstream of TGFβ, and have distinct transcriptional target genes (Felici et al, 2003; Furuhashi et al, 2001; Ju et al, 2006; Kim et al, 2005; Kretschmer et al, 2003; Kurisaki et al, 2001; Levy and Hill.

2005; Liu et al, 2003; Selvamurugan et al, 2004; Tang et al, 2003; ten Dijke and Hill,

2004; Uemura et al, 2005). For example, using Smad2- and Smad3-deficient mouse embryonic fibroblasts, it has been shown that TGFβ-mediated induction of matrix metalloproteinase-2 is selectively dependent upon Smad2, whereas induction of c-fos and 117 of Smad7 relies on Smad3 (Piek et al, 2001). Selvamurugan et al demonstrated a cooperative physical interaction between Runx2 and Smad2 proteins, but not between

Runx2 and Smad3, conferring the maximal collagenase-3 promoter activity induced by

TGFβ in rat osteoblastic cell line UMR 106-01 (Selvamurugan et al, 2004). Our previous results have shown that blocking DYNLRB1 expression disrupts Smad2-dependent, but not Smad3-dependent, transcriptional activation (Jin et al, in revision). However, Felici et al reported that TLP interacts predominantly TβRII, and over-expression of TLP interferes with Smad3–Smad4 complex formation and impairs Smad3-dependent transcriptional response (Felici et al, 2003). Smad3 has been demonstrated to be integral for transactivation of human Smad7 promoter and PAI-1 induction by TGFβ in many cells, but no compensation by the highly homologous Smad2 is observed in the Smad3 null fibroblasts disruption of such induction by Smad3 function cannot be compensated by Smad2 when a Smad3 specific inhibitor or dominant negative Smad3, or Smad3 null cells are employed (Datto et al, 1999; Frederick et al, 2004; Liu et al, 2006d; Piek et al,

2001; von Gersdorff et al, 2000).

In keeping with those in the field, results in this Chapter demonstrated that blocking DYNLRB2 expression specifically interferes with Smad3-dependent transactivation of human Smad7 promoter and PAI-1 induction by TGFβ, which could not be compensated by DYNLRB1, indicating a requirement of DYNLRB2 in specific

Smad3-dependent transcriptional regulation. However, the effect of blocking DYNLRB2 expression on reporters with artificial promoters or promoters from endogenous genes may not reflect its effect on the regulation of endogenous genes. Since the regulation of 118 endogenous genes may have substantially different requirements from the regulation of reporters with promoters from endogenous genes from a transiently transfected vector.

Gene expression (especially single gene expression) alone may not reflect the overall effect of TGFβ signaling on the cell (such as growth inhibition and differentiation).

Therefore, the physiological significance of this requirement of DYNLRB2 in Smad3 specific transcriptional regulation remains to be determined in global effect (such as growth inhibition) in DYNLRB2-knockout cells and animal models. But, many knockout mice targeting TGFβ signaling components die at an early stage during embryogenesis (Table 1) and make it unable to investigate the effects of a complete loss of function of these components in differentiated tissues, such as Smad2 and Smad4

(Weinstein et al., 1998; Yang et al., 1999). Thus, it would be better to make conditional knockout mouse with targeted loss of DYNLRB2 in skin, or ovary, to explore the function of DYNLRB2 in these tissues, since in this Chapter we have demonstrated a role for DYNLRB2 in Smad3-dependent TGFβ signaling in keratinocytes (HaCaT cells) and it has been demonstrated that ovarian follicle development requires Smad3 by knockout mouse model (Tomic et al, 2002 and 2004). Phenotype similar to Smad3-knockout mice or some phenotype overlap might be expected, such as ovarian follicle development defects or skeletal defects. An inducible knockdown approach might be employed to investigate the role of DYNLRB2 in cells and animal models, because of its several advantages. The wild-type and engineered cells or animals will be on an isogenic background, and they will not accumulate mutations or adjust expression levels of other genes that might compensate for the loss of DYNLRB2, which might occur when

DYNLRB2-null cells are used. Another advantage is the opportunity to compare the 119 effects of different levels of DYNLRB2 in the cell or animal model, by adjusting the level of inducing reagents (like Tetracycline), since different effects of TGFβ might require different levels of DYNLRB expression.

A prominent biological effect of TGFβ in epithelial cells is growth inhibition

(Yue and Mulder, 2001; Derynck and Zhang, 2003). Since DYNLRB1 and 2 are 77% identical in amino acid sequence, and our previous results have shown that DYNRB1 is partially required for mediating the growth inhibitory response to TGFβ (Jin et al, 2005), it might be expected that DYNLRB2 share this functional feature with DYNLRB1. Our preliminary results suggest that blockade of endogenous DYNLRB2 in MDCK cells with high passage numbers (between 25 and 30) after thawing from stock decreased the ability of TGFβ to inhibit DNA synthesis measured by [3H]-thymidine incorporation assay

(preliminary results in Appendix A), indicating that DYNRBL2 may be partially required for TGFβ-mediated inhibition of DNA synthesis, similar to DYNLRB1. But this effect was not observed in MDCK cells with low passage numbers (between 10 and 15) after thawing from stock (preliminary results in Appendix A). This is intriguing, and it is currently unknown how blocking DYNLRB2 expression exerts such different effects in

MDCK cells with high and low passage numbers, respectively. However, Liu et al reported similarly that TGFβ mediates activation of Smad2 primarily in early cultures (1 day) of primary rat hepatic stellate cells, but activates Smad3 primarily in late transdifferentiated cultures (7 day) of primary rat hepatic stellate cells (Liu et al, 2003).

While this could suggest that DYNLRB2 is involved in Smad3-dependent signaling related with growth inhibition, it is also possible that DYNLRB2 is involved one of other 120 signaling pathways that are involved in mediating this complex biological response. For example, TGFβ activation of the Smad-independent pathways is also partially required for negative growth control by TGFβ, such as TGFβ-mediated activation of Ras, ERKs and JNKs (Derynck and Zhang, 2003; Moustakas and Heldin, 2005; Mulder, 2000; ten

Dijke and Hill, 2004). However, the Smad-independent pathways, their interactions with each other, and their roles in TGF-β-mediated growth inhibitory effects are not well understood. Thus, it is clear that more studies are warranted to investigate the role of

DYNLRB2 in TGFβ-mediated growth inhibition in DYNLRB2 loss-of-function cells or cells with stable inducible expression of DYNLRB2 siRNAs, as discussed above.

Recent studies have demonstrated that endocytosis, particularly the clathrin- mediated pathway, plays a critical role in the early events of TGFβ signal transduction

(Anders et al, 1997; Di Guglielmo et al, 2003; Hayes et al, 2002). However, it is unclear how signaling components (like Smad2 and Smad3) move close to the nucleus before their translocation into the nucleus. Previous results from our lab and others have suggested that the dynein motor may target TGFβ signaling components for intracellular transport to specific subcellular compartments through specific dynein light chains (Ding and Mulder, 2004; Jin et al, 2005 and in revision; Machado et al, 2003; Tang et al, 2002).

For example, it has been demonstrated in the lab that DYNLRB1 interacts and is co- localized with Smad2 at early times after TGFβ treatment (Jin et al, in revision).

Blocking DYNLRB1 has been shown to significantly impair the translocation of Smad2 into the nucleus (Jin et al, in revision). Similarly, Tang et al have shown that TGFβ stimulates phosphorylation of cytoskeletal adaptor protein β-spectrin ELF and its 121 subsequent association and colocalization with Smad3 (Tang et al, 2003). They have also shown that β-spectrin ELF deficiency results in mislocalization of Smad3 and disruption of the TGFβ-dependent transcriptional response (Tang et al, 2003). These results suggest that β-spectrin ELF functions as an essential adaptor protein specific for Smad3 and facilitates its subcellular localization for optimal signal propagation. Here results in this

Chapter provided evidence for such a model that DYNLRB2 interacts preferentially with

Smad3, and TGFβ stimulates the phosphorylation of DYNLRB2 to activate its recruitment to the dynein motoris, thus potentially allowing its targeting Smad3 to dynein motor for intracellular transport and optimal signal propagation. Therefore, these results are keeping with those in the field, and provide more evidence and details for the distinct pathways and unique roles of Smad3-dependent TGFβ signaling. However, the lack of

DYNLRB2 antibodies prevented us from investigating the interaction between endogenous DYNLRB2 and dynein motor, as well as the interaction between endogenous

DYNLRB2 and Smad3, and the phosphorylation status of its endogenous proteins upon

TGFβ stimulation. Therefore, once DYNLRB2 antibodies become available, this model can be tested in the future by examining the interaction between the endogenous molecules, and the subcellular localization and nuclear translocation of Smad3, after

DYNLRB2 knockdown or preferentially in DYNLRB2 knockout cells.

Overall, the results in this Chapter demonstrate for the first time a distinct regulation of Smad3-dependent TGFβ signaling by the DYNLRB2 dynein light chain. 122

Chapter 4

Requirement of zebrafish dynein light chain zDYNLRB in TGFβ signaling in

zebrafish ovarian follicle cells

Dyneins are molecular motors that play a wide variety of important functions in the cells. It has been demonstrated that DYNLRB family dynein light chains are required in TGFβ signaling in mammalian cells. Zebrafish

(Danio rerio) DYNLRB is highly homologous to human DYNLRB1 and

DYNLRB2. To explore the function of the zDYNLRB, the regulation of zDYNLRB function by TGFβ was investigated, as well as its role in TGFβ- mediated transcriptional activation in zebrafish ovarian follicle cells (zOFCs). It has been demonstrated that zDYNLRB was rapidly phosphorylated after TGFβ stimulation. In addition, it has been shown that the TβRII kinase is required for this phosphorylation. It has also been demonstrated that TGFβ stimulated a rapid recruitment of the zDYNLRB to the dyenin motor. Further, the phosphorylation of zDYNLRB was demonstrated to be responsible for this recruitment. Lastly, it has been demonstrated that zDYNLRB knockdown by morpholino significantly inhibited TGFβ-mediated induction of TRE-Luc, 3TP-Lux and ARE-Lux in zOFCs, but not phTG5-Lux transcriptional activation. Collectively, experiments in this chapter characterized the regulation of zDYNLRB by TGFβ and indicated a role for zDYNLRB in TGFβ signaling in zebrafish ovarian follicle cells. 123 4.1 Introduction

Women, like other female mammals, are born with a finite reserve of follicles in the ovary, which will only decrease over the lifetime (Bristol-Gould et al, 2006; Zeleznik,

2004). They have a dynamic reproductive system with cyclic fluctuations in the secretion of extraovarian factors (mostly from the brain) and intraovarian factors, which regulate the development of ovarian follicles (Barrett et al, 2006; Duggavathi et al, 2005; Knight and Glister, 2003). Thus, women's reproductive health covers the whole lifetime and focuses on ovarian development and functions. Diseases involving abnormal ovarian follicle development include infertility, polycystic ovarian syndrome, premature ovarian failure, ovarian teratoma and ovarian cancer (Azziz et al, 2005; Blank et al, 2006;

D'Andrilli et al, 2004; Meskhi and Seif, 2006; Nelson et al, 2005). However, the mechanisms underlying normal and abnormal ovarian follicle development are not clear.

The structural and functional unit of the ovary is the follicle, which comprises the oocyte and the surrounding layer(s) of somatic cells (Knight and Glister, 2003).

Complex cell-to-cell interactions coordinate the ovarian follicle development, and disruption of such interactions can lead to failure of oocyte development (Knight and

Glister, 2003; Ge, 2005). In vertebrates, it has been established that the somatic cells of the follicle provide a proper yet dynamic microenvironment that supports and nurtures the appropriate development of the oocyte (Eppig et al, 2002; Knight and Glister, 2003;

Ge, 2005). However, it is not completely known yet whether one cell type, either the somatic cell or the germ cell, or both determine the overall rate of follicle development

(Erickson and Shimasaki, 2000; Peng et al, 2000; Eppig et al, 2002; Ge, 2005). Thus, a 124 more in-depth understanding of the regulation of ovarian follicle development is required for the development of new diagnostics and effective therapeutic strategies for the detection and management of diseases, such as premature ovarian failure, polycystic ovarian syndrome, ovarian cancer and infertility.

The multi-functional TGFβ family signaling pathways play important roles in a wide range of processes such as adult tissue homeostasis and organogenesis of many organ systems, including the ovary (Massague, 1998; Heldin et al, 1997; Knight and

Glister, 2003; Pangas and Matzuk, 2004). It has been demonstrated that various TGFβ sumperfamily members (TGFβ, Activin, Inhibin, GDF9, etc.) are involved in important aspects in follicle development, and TGFβ signaling intermediates (like Smad3) are required for ovarian follicle development (Tomic et al, 2002 and 2004; Kohli et al, 2003;

Knight and Glister, 2003; Pangas and Matzuk, 2004; Kohli et al, 2005). However, inconsistent findings on the involvement of TGFβ in ovarian follicle development have been reported in mammals and several species as well (Attia et al, 2000; Coskun and Lin,

1994; Feng et al, 1988; Juneja et al, 1996; Knight and Glister, 2003; Kohli et al, 2003;

Liu et al, 1999; Pangas and Matzuk, 2004; Roy, 1993; Singh, et a, 1993; Tsafrirri et al,

1989), suggesting that more studies are required to investigate the precise role of TGFβ in the ovary.

Cytoplasmic dyneins are molecular motors that play a wide variety of functions in the cells, including transport of various intracellular cargoes, such as mRNA, and organelles like endosomes (Hirokawa, 1998; Kamal and Goldstein, 2002; Mallik and

Gross, 2004; Vale, 2003; Vallee et al, 2004). The dynein motor complex is a large 125 multimeric complex. Three distinct families of DLCs (the DYNLL, DYNLT, and

DYNLRB) have been identified in mammalian cells (Pfister et al, 2005; Vale, 2003;

Williams et al, 2005; Wu et al, 2005). DLCs have been shown to interact with a number of cargoes to exert diverse functions (Kamal and Goldstein, 2002; Mallik and Gross,

2004; Vallee et al, 2004). Robl is a mutant Drosophila DYNLRB family DLC (Jin et al,

2005; Bowman et al, 1999). In addition to defects in intracellular transport, Drosophila mutants in this gene (like the roblz deletion mutants) show a female sterile phenotype, suggesting a role of this gene in the ovary (Bowman et al, 1999). DYNLRB1 was identified as a TGFβ receptor-interacting protein, and is also a DLC of the DYNLRB family (Tang et al, 2002; Jin et al, 2005). It appears to play an important role in Smad2- dependent as well as other TGFβ signaling in mammalian cells (Ding and Mulder, 2004;

Jin et al, 2005 and in revision; Tang et al, 2002). In addition, DYNLRB1 is located on a chromosome region (human chromosome 20q12-q13.11), which is frequently amplified in ovarian cancers (Tanner et al, 2000), and DYNLRB1 mutations have been identified in ovarian cancer patient tumor tissues, implicating a role for this gene in the ovary (Ding et al, 2005). The results from Chapter 3 demonstrate a requirement of DYNLRB2 for

Smad3-dependent TGFβ signaling in mammalian cells. Smad3 is an important mediator of the TGFβ signaling pathway. It has been demonstrated that Smad3 knockout mice have reduced fertility compared with wild-type mice, and the reduced fertility in Smad3 knockout mice is due to impaired ovarian follicle development, associated with altered expression of gene that control cell cycle progression, cell survival and cell differentiation (Tomic et al, 2002 and 2004), thus providing strong evidence that ovarian follicle development requires Smad3. Therefore, we hypothesized that the DYNLRB 126 family DLC might play an important role in TGFβ signaling in ovarian follicle cells, which may be required for normal ovarian follicle development.

Recent studies have demonstrated that zebrafish is an excellent model for the investigation of ovarian follicle development, due to its advantageous attributes, such as small body size, easy maintenance, short life cycle, fast development in vitro and large-scale genetic screening (Fishman, 2001; Wu et al,

2000; Pang and Ge, 2002a; Ge, 2005). Primary culture of zebrafish ovarian follicles and ovarian follicle cells are well established and used previously successfully in many studies (DiMuccio et al, 2005; Kohli et al, 2003 and 2005;

Pang and Ge, 1999; Pang and Ge, 2002a and b; Wu et al, 2000). Zebrafish

(Danio rerio) DYNLRB (termed zDYNLRB hereafter) shows 80% and 77% amino acid sequence identity to human DYNLRB1 and DYNLRB2, respectively.

To take advantage of this model, we want to test our hypothesis in zebrafish ovarian follicle cells that the function of zDYNLRB might be regulated by TGFβ, and zDYNLRB might play an important role in TGFβ signaling in zebrafish ovarian follicle cells. There is evidence that for the presence of the TGFβ signaling pathway major molecules in the fish ovary (Kohli et al, 2003 and 2005;

Calp et al, 2003; Dick et al, 2000; Hardie et al, 1998). Recent findings suggest that TGFβ may play a role in preventing premature oocyte maturation in zebrafish by down-regulating basal and human chorionic gonadotropin-induced 20β- hydroxysteroid dehydrogenase, luteinizing hormone receptor, and membrane progestin receptor-β mRNA levels (Kohli et al, 2003 and 2005). Results in this 127 chapter demonstrate that the function of zDYNLRB is regulated by TGFβ in zOFCs, and zDYNLRB is required for TGFβ-mediated induction of TRE-Luc,

3TP-Lux and ARE-Lux in zOFCs, implicating a role for zDYNLRB in zebrafish ovarian follicle development.

4.2 MATERIALS AND METHODS

Reagents--The anti-Flag M2 (F3165) and mouse IgG were from Sigma. The anti-DIC monoclonal Ab was from Chemicon (Temecula, CA). The mouse anti-Smad2/3 (Cat. #

610843) was from BD Biosciences Transduction Laboratories (Palo Alto, CA). Our rabbit anti- hDYNLRB1 antibody was prepared against the amino acids 27–43 of hDYNLRB1 (GIPIKSTMDNPTTTQYA), purified by Strategic BioSolutions (Newark,

DE) and have been used successfully previously (Jin et al, 2005). The rabbit IgG, mouse

IgG and protein A/G plus agarose were from Santa Cruz Biotech. 32P-orthophosphate

(NEX-053) and 3H-thymidine (NET-027X) were from Perkin Elmer (Boston, MA).

TGFβ was purchased from R & D Systems (Minneapolis, MN). The Fugene 6 transfection reagent was from Roche Applied Science (Indianapolis, IN). The

Lipofectamine™ 2000 transfection reagent was from Invitrogen (Carlsbad, CA). The

Dual-Luciferase Reporter Assay System (Cat. # E1960) was purchased from Promega

(Madison, MI).

Cell Culture-- Mv1Lu cells (CCL-64) were obtained from ATCC (Rockville, MD) and were grown in DMEM supplemented with 10% FBS. DR26 cells, a Mv1Lu derivative 128 lacking functional TRII (Laiho et al, 1990 and 1991), were kindly provided by Dr. Joan

Massague (Sloan-Kettering) and maintained as for the Mv1Lu cells. 293T cells were obtained from T-W. Wong (Bristol-Myers Squibb) and were maintained as for Mv1Lu cells. Cultures were routinely screened for mycoplasma using Hoechst staining.

Isolation of zebrafish Ovarian follicles and Primary Culture of the Follicle Cells were performed essentially as described previously (Pang and Ge, 2002a), except that the dispersed follicles were washed three times with medium 199 with 10% FBS and 100

U/ml penicillin/100 µg/ml streptomycin (Invitrogen, Cat. # 15140), and incubated in culture dishes with the same medium for two days, before changing to M199 with

10%FBS.

Extraction of total RNA from the cultured follicle cells and RT-PCR amplification of zDYNLRB: zOFCs were plated in 6-well plates the day before. After discarding the medium, 1 ml Trizol Reagent from Invitrogen (Cat. No. 15596-026) was added to each well and total RNA was extracted essentially according to the manufacturer’s instructions. Dissolved total RNA in DEPC-treated water was stored in an -80 °C freezer. RNA is reverse transcribed to make cDNA using Omniscript Reverse

Transcriptase (QIAGEN) and Superscript First-Strand Synthesis System for RT-PCR

(Invitrogen) with random hexamer primers. PCR amplification of zDYNLRB is performed with Forward primer (5’ GCC GAG GTG GAG GAG ACT AT 3’) and

Reverse primer (5’ TGT GGG ATT TTG AAT GAC GA 3’). Control is 18s rRNA with primers from Eurogentec (San Diego, CA). PCR conditions are as follows: 94°C 50’,

60°C 60’, 72°C 60’ for 36 cycles. 129 Construction of pCMV5-zDYNLRB-Flag plasmid: To prepare zDYNLRB-2-Flag, zDYNLRB was PCR amplified from the above cDNA produced from the cultured follicle cells, using primers containing additional suitable flanking restriction enzyme sites for

BglII (5’) and SalI (3’), and inserted into pCMV5-Flag (Sigma) after digestion with BglII and SalI restriction enzymes, respectively. The correct DNA sequences were confirmed by sequencing and alignment to ZFIN (GenBank Accession number: NM_201188).

Transient transfections, immunoprecipitation/blot, Westerns, and in vivo phosphorylation were performed essentially as described previously (Tang et al, 2002;

Jin et al, 2005; Chapter 2 and 3).

Morpholino oligos (MOs): Morpholino oligos (Ekker and Larson, 2001) with the following sequences (ATG morpholino oligo (ATG MO): 5’-TAA TAG TCT CCT CCA

CCT CGG CCA T-3’, 5-mismatch ATG morpholino oligo control (5-mismatch MO): 5’-

TAA TAc TCT CgT CCA gCT CcG CgA T-3’, 5’UTR morpholino oligo (UTR MO): 5’-

GAA GAC AAA CCG CTG TTT TCG TTG C-3’, splicing blocking morpholino oligo

(SB MO): TCT CTA ATG TAT CTT TAC CTG GTG C, and standard control ATG morpholino oligo (standard MO): 5’-CCT CTT ACC TCA GTT ACA ATT TAT A-3’) were ordered from Gene Tools, LLC (Philomath, OR). SB Mos are labeled with

Fluorescein.

Morpholino oligo delivery into zOFCs: Morpholino oligo deliverey into the primary zebrafish ovarian follicle cells by Endoporter (Gene Tools, LLC) were essentially performed according to the manufacturer’s instructions (Boyden ED and Dietrich WF. 130 2006; Choudhury et al, 2006; Masaki et al, 2005; Masaki et al, 2005; Summerton JE.

2005; Tyson-Capper and Europe-Finner. 2006).

Morpholino Microinjection in zebrafish embryos: Wild-type zebrafish embryos were obtained by natural mating. zDYNLRB ATG MO and SB MO were injected into the yolk sac of the one- to two-cell embryos. Four hr postinjection, damaged embryos were sorted out and the rest were allowed to grow at 28°C for further observation and photographing. Control injections were done with injection buffer.

Immunofluorescence staining analysis: For Smad2/3 Immunofluorescence staining experiments, zOFCs were fixed with 4% paraformaldehyde in phosphate-buffered saline for 20 min at room temperature, and permeabilized with 0.5% Triton X-100 in phosphate-buffered saline for 5 min. Subsequently, these cells were incubated with 10

µg/ml anti-Smad2/3 monoclonal Ab for 1 h. The bound primary antibodies were visualized with 2 µg/ml cy3 -conjugated goat anti-mouse IgG (red). Images were collected with a Leica TCS SP2 AOBS confocal microscope, or a Nikon Diaphot microscope with a Retiga 1300 CCD camera (BioVision Technologies, Inc, Exton, PA) running IPLab v3.6.3 software (Scanalytics, Inc, Fairfax, VA). DAPI staining was used to visualize the nucleus of individual cells.

Luciferase reporter assay: zOFCs were plated (1 to 3 split) in 24-well plates. Twenty- four hours after plating, the cells were transfected using FuGene 6 transfection reagent with 0.1 µg/well TRE-Luc (Renshaw et al, 1995; Zhang et al, 1998), 3TP-Lux (Wrana et al, 1992; Lagna et al, 1996), phTG5-Lux (Yue and Mulder, 2000; Kim et al, 1990), or 131 ARE-Lux (Chen et al, 1997; Yeo et al, 1999), and 0.1 µg /well pRL-SV (control to normalize transfection efficiencies), followed by delivery of MOs by Endoporter for 6 h,

12 h after transfection. Then the cells were washed with SF and incubated in this SF for

30 min, prior to an incubation of 24 h in the presence or absence of TGFβ (10ng/ml).

Then luciferase activity of the cell lysates was measured using the Dual-Luciferase

Reporter Assay System following the manufacturer’s instructions. All assays were performed in triplicate. Data are expressed as mean±SEM.

Growth assays: The TGFβ growth inhibitory responsiveness of the zOFCs was assessed by [3H] thymidine incorporation assays, performed as described (Jin et al, 2005;

Hartsough and Mulder, 1995).

4.3 Results

4.3.1 Cloning and expression detection of zDYNLRB

To test our hypothesis that the function of zDYNLRB might be regulated by

TGFβ, zDYNLRB was cloned and inserted into the pCMV5-Flag vector (Materials and

Methods). Blast search in the NCBI and Ensembl database showed that there is only one gene found so far for this family DLCs in zebrafish. But, it is quite possible that there may be more than one gene for this DLC family in zebrafish and such genes may be identified when the sequencing of zebrafish genome is finished. The DYNLRB DLC family displays a considerable degree of conservation across different Kingdoms and

Phyla (Tang et al, 2002; Koonin and Aravind, 2000; Jin et al, 2005). Sequence alignment 132 (Fig. 18) showed that overall zDYNLRB is 80% and 77% identical in amino acid sequence to human DYNLRB1 and DYNLRB2, respectively, and it is more similar to human DYNLRB2 in its middle region of the protein, while it displays more similarity to human DYNLRB1 in the protein’s two terminal regions.

Expression of zDYNLRB-Flag was confirmed by Western blot with an anti-

Flag Ab (data not shown). To determine whether zDYNLRB could be detected by the rabbit anti-DYNLRB1 antibody (DYNLRB1 Ab), which has been used successfully to detect human DYNLRB1 and the peptide sequence to make the antibody is only one amino acid different, as described previously (Jin et al, 2005), immunoprecipitation

(IP)/blot analyses were performed using an anti-Flag Ab as the IP Ab and DYNLRB1 Ab as the blot Ab, after transfection of zDYNLRB into 293T cells. As shown in Fig.19A, expression of transfected zDYNLRB-Flag was detectable using DYNLRB1 Ab (lane 2), while empty vector (EV) control was negative (lane 1), indicating that the DYNLRB1 Ab could detect exogenous zDYNLRB.

To determine whether DYNLRB1 Ab could also detect endogenous zDYNLRB,

Western blot analyses was performed on cell lysates from zOFCs. Primary zebrafish

OFC culture was adapted from methods established by Ge and colleagues (Pang and Ge,

2002a and b). As indicated in Fig. 19B, DYNLRB1 Ab recognized a single band of

11kDa (lane 1), whereas no band was detectable when IgG control was used (lane 2), indicating that DYNLRB1 Ab can also detect endogenous zDYNLRB in zOFCs by western blot. 133

Formatted Alignments

10 20 30 10 20 30 km-2phDYNLRB2 M A E V E E T L K R I Q S H K G V I G T M V V N A E G I P I km-2p M A E V E E T L K R I Q S H K G V I G T M V V N A E G I P I hkm23-1p M A E V E E T L K R L Q S Q K G V Q G I I V V N T E G I P I hkm23-1phDYNLRB1 M A E V E E T L K R L Q S Q K G V Q G I I V V N T E G I P I zkm23p M A E V E E T I K R I Q S Q K G V Q G I I I V N A E G I P I zkm23pzDYNLRB M A E V E E T I K R I Q S Q K G V Q G I I I V N A E G I P I M A E V E E T L K R I Q S Q K G V Q G I I V V N A E G I P I M A E V E E T L K R I Q S Q K G V Q G I I V V N A E G I P I

40 50 60 40 50 60 km-2p R T T L D N S T T V Q Y A G L L H H L T M K A K S T V R D I km-2phDYNLRB2 R T T L D N S T T V Q Y A G L L H H L T M K A K S T V R D I hkm23-1p K S T M D N P T T T Q Y A S L M H S F I L K A R S T V R D I hkm23-1phDYNLRB1 K S T M D N P T T T Q Y A S L M H S F I L K A R S T V R D I zkm23p K S T L D N T S T V Q Y A A N I H Q L L M K A R G I V R D I zkm23pzDYNLRB K S T L D N T S T V Q Y A A N I H Q L L M K A R G I V R D I K S T L D N . T T V Q Y A . L . H L . M K A R S T V R D I K S T L D N . T T V Q Y A . L . H L . M K A R S T V R D I

70 80 90 70 80 90 km-2p D P Q N D L T F L R I R S K K H E I M V A P D K E Y L L I V km-2phDYNLRB2 D P Q N D L T F L R I R S K K H E I M V A P D K E Y L L I V hkm23-1p D P Q N D L T F L R I R S K K N E I M V A P D K D Y F L I V hkm23-1phDYNLRB1 D P Q N D L T F L R I R S K K N E I M V A P D K D Y F L I V zkm23p D P Q N D L T F L R V R S K K N E I M I A P D K D Y F L I V zkm23pzDYNLRB D P Q N D L T F L R V R S K K N E I M I A P D K D Y F L I V D P Q N D L T F L R I R S K K N E I M V A P D K D Y F L I V D P Q N D L T F L R I R S K K N E I M V A P D K D Y F L I V

100 110 120 100 110 120 km-2p I Q N P C E * km-2phDYNLRB2 I Q N P C E * hkm23-1p I Q N P T E * hkm23-1phDYNLRB1 I Q N P T E * zkm23p I Q N P T E * zkm23pzDYNLRB I Q N P T E * I Q N P T E I Q N P T E

Fig. 18. Amino acid sequence alignment of hDYNLRB1, hDYNLRB2 and zDYNLRB. hDYNLRB1, hDYNLRB2 and zDYNLRB amino acid sequences were obtained from NCBI database (Accession number: NP_054902, NP_570967 and AAH46084, respectively) and aligned using the MacVector software. 134

A B

293T zOFC pCMV5-EV + - kDa kDa zDYNLRB - + zDYNLRB 14 14 zDYNLRB 6 1 2 1 2 DYNLRB1 IP Ab: Flag Blot Ab: IgG Blot Ab: DYNLRB1

Fig. 19. Detection of zebrafish DYNLRB. A: 293T cells were transiently transfected with zDYNLRB-Flag (Lane 2) or EV (Lane 1). Thirty-two hours after transfection, cells were lysed, and IP/blot analyses were performed. The lysates were IP'd with an anti-Flag Ab. IP’d lysates were separated by 4-12% NuPAGE, transferred to a PVDF membrane, and blotted with the rabbit polyclonal DYNLRB1. B: Western blot analysis of endogeous zDYNLRB in zOFCs using DYNLRB1. zOFC lysates were separated by 4- 12% NuPAGE, and transferred to a PVDF membrane. Western blot analyses were performed using DYNLRB1 Ab (Lane 1), with rabbit IgG as a control (Lane 2). 135 4.3.2 Primary zebrafish ovarian follicle cells (OFCs) are TGFβ responsive

Culture of zebrafish primary ovarian follicle cells were established by Ge and colleagues (Pang and Ge, 2002a and b). Most of the cells present in the zOFC culture are the two epithelial cells with distinct morphology (the thecal cells and the granulosa cells).

The primary zOFC culture is performed as described previously (Pang and Ge, 2002a).

Since our study relies upon the responsiveness of the cells to TGFβ treatment, thymidine incorporation analyses were performed to determine whether zOFCs are responsive to the growth inhibitory effects of TGFβ. As shown in Fig. 20A TGFβ treatment for 24 h induced a 50% growth inhibition of the zOFCs. This result is consistent with a recent report indicating TGFβ significantly inhibited both gonadotropin- and 17α, 20β- dihydroxyprogesterone-induced oocyte maturation in a dose- and time-dependent manner

(Kohli et al, 2003).

To further confirm the responsiveness of the zOFCs to TGFβ treatment, immunofluorescence staining experiments were performed to determine whether endogenous Smad2/3 proteins is translocated from the cytoplasm into the nucleus, since

Smad2/3 are the TGFβ major signaling components and have been cloned from the zebrafish ovary (Kohli et al, 2003; Dick et al, 2000). Results from Fig. 20B show that

TGFβ treatment for 15 min stimulates the cytoplasmic Smad2/3 (punctate cytoplasmic staining in the absence of TGFβ) go into the nucleus (condensed nuclear staining in the presence of TGFβ, counter-stained by DAPI). These results are in agreement with the recent reports that cloned TGFβ, TGFβ receptors, Smad proteins and other signaling

137 intermediates as well from the zOFCs (Calp et al, 2003; Dick et al, 2000; Hardie et al,

1998; Kohli et al, 2003). Thus, these experiments clearly showed that primary zOFCs are responsive to TGFβ treatment.

4.3.3 zDYNLRB is phosphorylated after TGFβ receptor activation

TGFβ receptors have serine/threonine kinase activity and can phosphorylate a number of intracellular proteins to initiate and propagate various TGFβ signaling events and responses (Shi and Massague, 2003; Attisano and Wrana, 2002; Yue and Mulder,

2001). The current model of the canonical signal initiation for the TGFβ cytokine is to signal through cell surface complexes of TβRI and TβRII (Shi and Massague, 2003;

Attisano and Wrana, 2002; Yue and Mulder, 2001). Ligand binding allows the formation of a stable TβRII/RI receptor complex and conformational change, allowing phosphorylation and activation of TβRI by the TβRII kinase, leading to phosphorylation of intracellular proteins such as Smad proteins, ERKs, and JNKs (Shi and Massague,

2003; Attisano and Wrana, 2002; Yue and Mulder, 2001). We have shown previously that DYNLRB1 is phosphorylated on serine residues upon TGFβ receptor activation

(Tang et al, 2002; Jin et al, 2005). Thus, if zDYNLRB is a component of the TGFβ signaling network, it might be expected that zDYNLRB be phosphorylated upon TGFβ receptor activation as a mechanism for its activation.

To determine whether zDYNLRB is phosphorylated upon activation of TGFβ receptors, in vivo phosphorylation assays were performed after transient transfection of zDYNLRB-Flag in 293T cells, together with various TGFβ receptor constructs or empty 138 vector (as indicated), in the absence or presence of TGFβ (as indicated). As shown in

Fig. 21A, zDYNLRB was not constitutively phosphorylated (lane 1, top panel), and co- expression of zDYNLRB with TGFβ receptors for only a limited time (24 h) in the absence of TGFβ didn’t induce its phosphorylation (lane 2, top panel). However, TGFβ treatment for 15 min after co-expression of TGFβ receptors with zDYNLRB resulted in significant zDYNLRB phosphorylation (lane 3, top panel). Furthermore, this phosphorylation of zDYNLRB was completely blocked upon expression of a kinase- deficient form of TβRII (TβKNRII) (lane 4, top panel). No detectable phosphorylation of zDYNLRB (lane 5, top panel) was observed after expression of a constitutively active

TβRI (TβRI-T204D), which has been shown to phosphorylate some TGFβ signaling intermediates, such as Smad2 (Barrios-Rodiles et al, 2005). The results of no detectable phosphorylation for zDYNLRB in the presence of co-expression of both TβRI-T204D and TβKNRII (lane 6, top panel) further indicated that zDYNLRB phosphorylation required the kinase activity of TβRII, but not constitutively active TβRI. The IgG control

(lane 7, top panel) indicates that the band noted is specific for zDYNLRB. Equal expression of zDYNLRB was confirmed by blotting with anti-Flag (bottom panel), after transferring the same gel to a Polyvinylidene Fluoride (PVDF) membrane. Thus, the results in Fig. 21A demonstrate not only that zDYNLRB is phosphorylated upon treatment of TGFβ, but also that the kinase activity of TβRII, but not a constitutively active form of TβRI, is required for zDYNLRB phosphorylation.

To determine whether TGFβ stimulation leads to zDYNLRB phosphorylation in primary zOFCs, in vivo phosphorylation assays were performed in zOFCs, after transient

140 co-expression of zDYNLRB with either TβRII (Fig. 21B, lanes 2-4 and 6), or TβKNRII

(Fig. 21B, lane 5), or EV (Fig. 21B, lane 1). As shown in Fig. 21B, ligand activation of

TGFβ receptors for 5 min resulted in a low level of zDYNLRB phosphorylation (lane 1).

However, ligand activation of exogenous TGFβ receptors in zOFCs resulted in a much higher level of zDYNLRB phosphorylation (lanes 3-4). Furthermore, this phosphorylation of zDYNLRB was significantly blocked upon expression of the

TβKNRII in the presence of TGFβ treatment for 15 min (lane 5 versus lane 4). The IgG control (lane 4, top panel) indicates that the band noted is specific for zDYNLRB. Thus, similar to the results in the 293T cells, zDYNLRB was phosphorylated upon TGFβ treatment in zOFCs and its phosphorylation required the kinase activity of TβRII.

4.3.4 TGFβ stimulates the interaction between zDYNLRB and DIC, which requires the TβRII kinase

As mentioned earlier, zDYNLRB is a member of the DYNLRB/robl/LC7 family of DLCs (Jin et al, 2005; Susalka et al, 2002; Tang et al, 2002). It is thought that

DLCs may be important for specifying the nature of the cargo to be transported by the motor (Mallik and Gross, 2004; Kamal and Goldstein, 2002). Therefore, extracellular factors (such as growth factors, cytokines, etc) may likely select particular DLCs to specify the cargo for the dynein motor to transport, thus making it necessary for their recruitment to the dynein motor in specific cellular contexts. It has been shown that

TGFβ receptor activation induces rapid recruitment of the DYNLRB1 and 2 to the dynein motor through DIC (Tang et al, 2002; Chapter 2 and 3). Similarly, a more recent report has indicated that BMPR-II receptor activation may trigger its association with the dynein

142 motor complex (Machado et al, 2003). Accordingly, it was of interest to determine whether TGFβ could stimulate the recruitment of the zDYNLRB DLC to the dynein motor complex.

To determine whether TGFβ could stimulate the interaction between the zDYNLRB DLC and the DIC, IP/blot analyses were performed using anti-DIC as the IP antibody and anti-FLAG as the blotting antibody after transient transfection of zDYNLRB-Flag in the mink lung epithelial cells (Mv1Lu). These cells are highly responsive to TGFβ, and are frequently used in studies of TGFβ signaling (Jin et al,

2005; Tang et al, 2002; Yoon et al, 2005; Dore et al, 1998; Yingling et al, 1997; Nakao et al, 1997; Feng and Derynck, 1997; Wells et al, 1997; Wang et al, 1996). As shown in

Fig.22A, TGFβ stimulated the recruitment of zDYNLRB to the DIC (lanes 3 and 4).

Expression of EV only (lane 1) and the IgG control (lane 5) indicated that the interaction noted is specific for zDYNLRB. Similar results were obtained for 293T cells, after co- expression of TβRI-V5, TβRII-HA, and zDYNLRB-Flag in the presence and absence of

TGFβ (data not shown). Thus, similar to human DYNLRB1 and 2, zDYNLRB can also be recruited to the dynein motor after TGFβ stimulation.

Next, similar IP/blot experiments were performed to determine whether TGFβ could also regulate the recruitment of endogenous zDYNLRB to the dynein motor in primary zOFCs expressing endogenous TGFβ receptors. To achieve this, DYNLRB1 Ab was as used as the blot Ab. Western blot experiments (Fig. 22 Inset) showed that the

DIC antibody we used before also worked well in zOFCs. As shown in Fig.22B, there

144

zOFC TGFβ (min) 0 5 10 15 20 30 15 kDa 14 zDYNLRB 6 1 2 3 4 5 6 7 IP Ab: DIC IgG

Blot Ab: DYNLRB1

Fig. 22B. The recruitment of endogenous zDYNLRB to the DIC in zOFCs is stimulated by TGFβ. zOFCs were incubated in SF medium for 60 min prior to addition of TGFβ (10 ng/ml) for the indicated times. Cell lysates were subjected to IP/blot analyses using an anti-DIC Ab or IgG control as the IP Ab and DYNLRB1Ab as the blot Ab. 145 was no interaction between the zDYNLRB DLC and the DIC in the absence of TGFβ

(lane 1). However, TGFβ induced a rapid recruitment of zDYNLRB to the DIC, occurring as early as 5 min after TGFβ treatment (lanes 2-6). The maximal increase in this interaction was reached after 10 min of TGFβ treatment (lane 2) and appeared to remain relatively constant until at least 30 min after TGFβ treatment (lane 6). The IgG control was negative (lane 7, top panel). To further confirm the zDYNLRB-DIC interaction and its regulation by TGFβ, IP/blot analyses of the reverse direction using the anti-DIC as the blotting Ab (Fig. 22C) show a rapid interaction between DIC and zDYNLRB after TGFβ stimulation. Collectively, these data demonstrated that TGFβ induced a rapid recruitment of zDYNLRB to the dynein motor via DIC, and that this

TGFβ–mediated rapid zDYNLRB recruitment also occurred in primary ovarian follicle cells with endogenous TGFβ receptors.

It has been demonstrated previously that the kinase activity of TβRII is required for the interaction between the human DYNLRB family DLC and the DIC (Tang et al,

2002; Chapter 2 and 3). Since it has also been demonstrated here that zDYNLRB can also be phosphorylated and that the kinase activity of TβRII is required for this phosphorylation, it would be of interest to determine whether the recruitment of zDYNLRB to the DIC require TβRII kinase activity. In order to investigate this possibility, IP/blot analyses similar to those in Fig. 22A were performed, after transient expression of zDYNLRB in Mv1Lu cells and DR26 cells (a Mv1Lu mutant derivative cell line lacking functional TβRII) (Laiho et al, 1990 and 1991). As shown in Fig.22D,

TGFβ stimulated recruitment of zDYNLRB to the DIC in Mv1Lu cells expressing

148 endogenous wild type Tβ RII and RI (lanes 1-3). However, in DR26 cells, the recruitment of zDYNLRB to the DIC after TGFβ treatment was barely detectable (lanes

4-6). No specific band was detectable in the IgG control. Equal expression and loading of DIC was confirmed (bottom panel). Thus, the TβRII kinase is required for the recruitment of zDYNLRB to the rest of dynein motor complex.

4.3.5 zDYNLRB specific morpholinos knock down zDYNLRB expression

The next to ask now is whether zDYNLRB is involved in TGFβ signaling in zOFCs. We chose a knockdown strategy to block zDYNLRB expression with morpholino phosphorodiamidate oligonucleotides (MOs), which block translation from target mRNA or blocking target mRNA splicing (Ekker and Larson, 2001; Nasevicius and Ekker, 2000). For MOs are very stable molecules and can persist over seven days after the delivery, because they are not degraded in cells. They are very effective to decrease gene expression even to levels undetectable by Western blot. Translation- blocking MOs was designed. To deliver MOs into cells, Endo-Porter was used, which is excellent for delivering MOs into cells (Boyden and Dietrich, 2006; Choudhury et al,

2006; Masaki et al, 2005; Masaki et al, 2005; Tyson-Capper and Europe-Finner, 2006).

It is a novel peptide explicitly designed to deliver substances into the cytosol of cells by an endocytosis-mediated process that avoids damaging the plasma membrane of the cell, which prevents loss of vital cell contents and toxicity (Summerton, 2005).

First, two zDYNLRB specific MOs were designed targeting the ATG start codon region and the 5’ untranslated region of the zDYNLRB mRNA, and their 149 translation blocking effect was tested in 293T cells and zOFCs. As shown in Fig. 23A, the zDYNLRB ATG MOs specifically knocked down exogenous zDYNLRB expression in 293T cells (top panel, Lanes 1-3), even at a concentration of 0.25µM, although a complete blocking effect was not achieved until a concentration of 4.0 µM zDYNLRB

ATG MOs was employed. However, no such effect was observed for the two control

MOs (top panel, lanes 4 and 5: standard control and zDYNLRB ATG MO 5-mismatch control, respectively) at1.0 µM, or the zDYNLRB UTR MOs (top panel, lane 6). Equal expression and loading of DIC was confirmed (bottom panel). The lack of effect of the zDYNLRB UTR morpholino oligo to knock down expression from the pCMV5- zDYNLRB-Flag plasmid is due to the reason that the target sequence for the zDYNLRB

UTR morpholino oligo was not constructed into the pCMV5-zDYNLRB-Flag plasmid.

Thus, the translation blocking effect of the zDYNLRB ATG MOs was dose-dependent and specific for zDYNLRB. The translation blocking effect of these MOs were then tested in the primary zOFCs. Results from Fig.23B showed that both zDYNLRB ATG

MOs and UTR MOs, but not the two control MOs, blocked endogenous zDYNLRB expression in zOFCs in a dose-dependent manner, with an optimal blocking concentration between 2.0 µM and 4.0 µM.

4.3.6 zDYNLRB knockdown interferes with TRE-Luc, 3TP-Lux and ARE-Lux induction by TGFβ, but not phTG5-Lux transcription activation

The TGFβ superfamily ligands were identified mainly through their roles in development, and they regulate a variety of processes such as cell proliferation, differentiation and migration, which are largely achieved through ligand-induced 150

293T zDYNLRB ATG MO (mM) .25 1.0 4.0 - - - 5-mismatch ATG MO (mM) - - - 1.0 - - standard MO (mM) - - - - 1.0 - zDYNLRB UTR MO (mM) - - - - - 1.0 kDa zDYNLRB-Flag(2 mg) + + + + + + zDYNLRB 14 6 1 2 3 4 5 6 Blot Ab: Flag 98 DIC 62 1 2 3 4 5 6 Blot Ab: DIC

Fig. 23A. The zDYNLRB MOs specifically knock down exogenous zDYNLRB expression in 293T cells. 293T cells were plated in 6-well plates, and twenty-four hours later, the cells were transiently transfected with 2µg zDYNLRB-Flag using FuGene 6 transfection reagent. Six hours after transfection, MOs were delivered to the cells by Endo-Porter at 0.25, 1.0 or 4.0 µM (as indicated). Forty-eight hours after transfection, cells are lysed. Top panel, blockade of zDYNLRB-Flag expression was analyzed by Western blotting with an anti-Flag M2 Ab (1:1000) on the lysates. Bottom panel, equal loading was confirmed by blotting with an anti-DIC Ab (1:1000).

152 transcription regulation of target genes (Attisano and Wrana, 2002; Derynck and Zhang.

2003; Roberts and Wakefield, 2003; Yue and Mulder, 2001). Now, with zDYNLRB specific MOs to knock down zDYNLRB expression, we determined whether zDYNLRB is required for transcriptional regulation by TGFβ in zOFCs. The effect of blocking zDYNLRB expression was determined on the induction of TRE-Luc by TGFβ. TREs are

12-O-tetradecanyl-13-acetate-responsive gene promoter elements, and are involved in the transcriptional responses of many TGFβ target genes (Kim et al, 1990; Zhang et al,

1998). TRE-Luc is a synthetic reporter commonly used in the TGFβ field, which contains four tandem AP1-binding sites from the collagenase I promoter (Renshaw et al,

1995; Zhang et al, 1998). As shown in Fig.24A, 24 h TGFβ treatment stimulated TRE-

Luc about 4 fold. Blocking zDYNLRB expression with zDYNLRB ATG MOs and UTR

MOs both significantly inhibited TRE-Luc induction by TGFβ, and dose-dependently, but not the lower concentration (0.5 µM), neither the two MOs controls at any of the three concentrations used. This result suggested that zDYNLRB might be involved in at least some TGFβ signaling events in zOFCs.

Next, the effect of zDYNLRB knockdown was determined with another commonly used reporter gene containing a more natural gene regulatory region, 3TP-

Lux, which is a chimeric reporter containing upstream regions from both the human plasminogen activator inhibitor 1 and the human collagenase gene (Wrana et al, 1992;

Lagna, 1996). As shown in Fig.24B, 3TP-Lux normally is induced about 8 fold by

TGFβ, and both zDYNLRB ATG MOs and UTR MOs at the concentration of 2.0 µM 153

zOFCs

1600016000 TGFβ(-) TGFβ(+) 14000 14000units 1200012000 10000

10000 luciferase

TRE-Luc 80008000

6000 Relative 6000 40004000 20002000 00 zDYNLRB ATG MO (mM)10.5 2 2.0 3 8.0 4 - 5 - 6 - 7 - 8 - 9- 10- 5-mismatch MO (mM) - - - 0.5 2.0 8.0 - - - - zDYNLRB UTR MO (mM) ------0.5 2.0 8.0 - standard MO (mM) ------2.0

Fig. 24A. MOs knocking down zDYNLRB expression markedly inhibited TRE-Luc induction by TGFβ. zOFCs were plated (1 to 3 split) in 24-well plates. Twenty-four hours after plating, the cells were transfected using FuGene 6 transfection reagent with 0.1 µg/well TRE-Luc, and 0.1 µg /well pRL-SV, followed by delivery of MOs with increasing concentrations (0.5 µM, 2.0 µM, 8.0 µM) and TGFβ treatment as described in Materials and Methods). Then luciferase assays were formed. Transfection efficiencies were normalized by renilla luciferase. All assays were performed in triplicate. Data are expressed as mean±SEM. 154

200000 zOFCs TGFβ(-) 150000150000

TGFβ(+) units

100000Lux 100000

luciferase luciferase 3TP-

5000050000 Relative Relative

00 standard MO 21 uM 2 - 3 - 4 - zDYNLRB ATG MO - 2 uM - - 5-mismatch MO - - 2 uM - zDYNLRB UTR MO - - - 2 uM

Fig. 24B. Blocking zDYNLRB expression also inhibited 3TP-Lux induction by TGFβ. Similar experiments were performed as in Fig. 24A, except using 3TP-Lux as the reporter gene and employing MOs (2.0 µM). 155 significantly repressed 3TP-Lux induction by TGFβ, further suggesting the role of zDYNLRB in TGFβ-mediated signaling in zOFCs.

Then the zDYNLRB knockdown effect was determined on a reporter gene with a natural promoter containing AP1-binding site, the phTG5-Lux (Yue and Mulder, 2000;

Kim et al, 1990), which contains a 450-base pair fragment of the human TGFβ1 gene promoter, since it has been reported that TGFβ is expressed in ovarian follicle cells (Calp et al, 2003; Kohli et al, 2003 and 2005) and autocrine production regulation of TGFβ is interesting area of research (Yue and Mulder, 2000). This reporter has been used in several published studies (Yue and Mulder, 2000; Liu et al, 2006a and b). However, as shown in Fig.24C, phTG5-Lux normally is only very weakly induced in zOFCs by 24 h

TGFβ stimulation, and both zDYNLRB ATG MOs and UTR MOs at the concentration of

2.0 µM did not have any effect on phTG5-Lux induction by TGFβ, nor on the basal level of phTG5-Lux transcription. But, the already high basal level the phTG5-Lux transcription suggests constitutive transcription of this gene in the zOFCs. These results suggest zDYNLRB might not be involved in phTG5-Lux transcription regulation in zOFCs.

Since it has been demonstrated that zebrafish oocyte maturation are enhanced by Activin-A and B (Pang and Ge, 1999 and 2002; Wu et al, 2000), whose signaling is also mediated by Smad2/3, similar to that of TGFβ. ARE-lux contains DNA sequences derived from the Xenopus Mix.2 gene promoter that convey TGFβ-Activin responsiveness through complexes of Smad2 and FAST-1, and ARE-Lux induction has been shown to be Smad2-dependent (Chen et al, 1997; Yeo et al, 1999). DYNLRB1, one 156

zOFCs TGFβ (-) 20000020000 TGFβ (+)

units 15000015000

Lux

-

100000luciferase 10000

phTG

50000Relative 5000

00 standard MO 2 uM1 -2 3- -4 zDYNLRB ATG MO - 2 uM - - ATG 5-mismatch MO - - 2 uM - zDYNLRB UTR MO - - - 2 uM

Fig. 24C. zDYNLRB expression knocking down did not interfere with phTG5- Lux transcription regulation. Similar experiments were performed as in Fig. 24B, except using phTG5-Lux as the reporter gene. 157 of zDYNLRB homologues in humans, has been demonstrated to be required for Smad2- dependent TGFβ signaling in the lab (Jin et al, in revision). Thus, the effect of zDYNLRB knockdown was examined on this widely used and more specific ARE-lux reporter. As shown in Fig.24D, TGFβ treatment for 24 h normally induced ARE-lux about 3 fold, and both zDYNLRB specific MOs at the concentration of 2.0 µM significantly impaired ARE-Lux induction by TGFβ, indicating a role for zDYNLRB in mediating TGFβ-Activin responsiveness to activate target genes transcription in zOFCs.

4.4 Discussion

Previously we identified a novel protein, DYNLRB1, which appears to play an important role in Smad2-dependent as well as other TGFβ signaling in mammalian cells

(Ding and Mulder, 2004; Jin et al, 2005 and in revision; Tang et al, 2002), and the results in Chapter 3 demonstrated a requirement of DYNLRB2 for Smad3-dependent TGFβ signaling in mammalian cells. Here we tested our hypothesis that the function of zDYNLRB might be regulated by TGFβ in zebrafish ovarian follicle cells, and zDYNLRB might regulate some TGFβ signaling events in such cells.

Results in this Chapter have shown that zDYNLRB phosphorylation is regulated by TGFβ in both 293T cells and the primary zOFCs, and this event requires the kinase activity of TβRII. It has also been shown that zDYNLRB interacts with the DIC to be recuited to the dynein complex, and zDYNLRB phosphorylation is required for this event to occur, suggesting that non-phosphorylated zDYNLRB is in an inactive state until its phosphorylation and activation to bind to the dynein complex and cargoes as well. 158

zOFCs 20000

units TGFβ (-)

15000 TGFβ (+) Lux

10000

luciferase luciferase ARE-

5000 Relative Relative 0 1 2 3 4 standard MO 2 uM - - - zDYNLRB ATG MO - 2 uM - - ATG 5-mismatch MO - - 2 uM - zDYNLRB UTR MO - - - 2 uM

Fig. 24D: Blocking zDYNLRB expression significantly repressed ARE-Lux induction by TGFβ. Left panel: similar experiments were performed as in Fig. 24A, except using ARE-Lux (co-transfected with 0.1 µg/well Fast-1plasmid) as the reporter gene and employing MOs (2.0 µM). Then luciferase activity was measured as described above. 159 These data are in agreement with results for hDYNLRB1 and 2 (Tang et al, 2002; Jin et al, 2005; Ilangovan et al, 2005; Chapter 2 and 3) and Tctex-1, a DLC interacting with

BMP receptors (Machado et al, 2003). zDYNLRB is very similar to human DYNLRB1 and 2, and overall displays 80% and 77% amino acid sequence identity to human

DYNLRB1 and 2, respectively, but it is more similar to DYNLRB2 in the middle region of the protein (Jin et al, 2005). The recent determination of the NMR and Crystal structures of human DYNLRB1 (Ilangovan et al, 2005; Liu et al, 2006c; Song et al,

2005b) identified a flexible (hence likely adaptable) deep inter-monomer cleft, which provides an ideal potential binding site for an array of structurally diverse partners, such as TGFβ receptors and Smads that DYNLRB family DLCs might bind. However, the location of residues that differ between zDYNLRB and hDYNLRB were suggested not to be localized in the above identified potential binding site for cargoes, implicating that residues that differ between zDYNLRB and hDYNLRB1 might not alter either DIC or cargo binding, which is supported by the results here in this Chapter for its interaction with DIC. With the structures now available, we have the resources to further dissect the structural basis for the function of zDYNLRB and its regulation mechanism(s).

It has also been demonstrated in this Chapter that blocking zDYNLRB expression with MOs resulted in significant inhibition of some common TGFβ signaling reporter genes’ transcriptional activation in zOFCs: TRE-Luc, 3TP-Lux and ARE-Lux, but not phTG5-Lux, suggesting that zDYNLRB is required for some TGFβ signaling events leading to transcriptional responses in the zebrafish ovary. However, a major limitation of this part of the thesis is the lack of physiological relevance, although 160 mutations have been identified in DYNLRB1 (one of zDYNLRB homologues in human) in human ovarian cancer patient tumor tissues, implicating a role for this gene in the ovary (Ding et al, 2005). In agreement with this, genetic evidence is provided to suggest a role for this gene in the ovary by the Drosophila homologue deletion mutants (roblz), which exhibit a female sterile phenotype (Bowman et al, 1999). Therefore, future experiments need to address the physiological significance of zDYNLRB function in the primary ovarian follicles by examining the effect of disrupting zDYNLRB function on

TGFβ targets in zebrafish ovarian follicle cells, which are recently identified and inhibit zebrafish oocyte maturation (Kohli et al, 2005). Or the function of zDYNLRB may be directly investigated on the ovarian follicle development and fertility of the mutant zebrafish with zDYNLRB function disrupted, in comparison with wild-type control.

Mutant zebrafish targeted at zDYNLRB may be made by TILLING (for Targeting

Induced Local Lesions in Genomes) (Amsterdam et al, 2006; Berghmans et al, 2005;

Wienholds et al, 2003). Then the two proposed experiments will be performed to determine the role of zDYNLRB in vivo on ovarian follicle development and zebrafish fertility. Its phenotype might be expected similar to or in overlap with some of the TGFβ signaling component mutants (Table 2).

Recently, it has been reported that TGFβ may acts upon multiple targets to inhibit zebrafish oocyte maturation in zebrafish (Kohli et al, 2005). However, Activin-A and Activin–B are reported to enhance zebrafish oocyte maturation (Pang and Ge, 1999 and 2002a; Wu et al, 2000), although TGFβ and Activin act through a similar Smad signaling pathway (Knight and Glister, 2003; Pangas and Matzuk, 2004; Massague, 161 Table 3. TGFβ family signaling pathway component mutants with ovary development defects

Genes Mutants Species Phenotypes References

TGFβ1 null mice viable and TGF 1 β knock mouse show severely impaired (ligand) Ingman et al, 2006 out reproductive capacity and almost complete infertility

BMP15 Female subfertility, decreased knock mouse Yan et al, 2001 (ligand) out ovulation and fertilization rates.

Complete female infertility; GDF9 knock Dong et al, 1996 mouse early block in folliculogenesis out Elvin et al, 1999 (ligand) at primary stage

Male infertility, female MIS/AMH knock Behringer et al, 1994 mouse premature loss of primordial out Durlinger et al, 1999 (ligand) follicles Coerver et al, 1996 Female infertility, secondary Inhibinα Matzuk et al, 1996 knock mouse infertility in males; granulose and (ligand) out sertoli cell and adrenal tumors Drummond et al, 2004

Activin Female infertility due to knock receptor mouse folliculogenesis defect; delayed Matzuk et al, 1995 out type II fertility in males; small gonads; (receptor) 25% of mice die at birth

Smad3 Slowed follicle growth, defective Tomic et al. 2002 knock (Transcription mouse folliculogenesis, degenerated and 2004 factor) out oocytes, and reduced fertility

Robl cDNA-rescued females show Bowman et al, 1999 roblz/null Drosophila (dynein fertility defect, fully rescued by light chain) genomic construct containing Robl Pentraxin 3 (Ptx3) knock Females display significantly mouse Varani et al, 2002 (GDF9 out reduced fertility target gene) 162 1998). Thus, TGFβ and Activin may have opposing effects on zebrafish oocyte maturation. Activin and TGFβ have also been reported to exert differential effects of on mouse and bovine ovarian follicle development as well (Liu et al, 1999; Hutchinson et al,

1987). However, currently, the underlying mechanisms are not clear for their opposing effects on ovarian follicle development. Recent structural studies showing that the

Activin and TGFβ ligands use quite distinct receptors for its signaling transmission rule out the possibility for the competition of the receptors (Shi and Massague, 2003; Hart et al, 2002; Guimond et al, 2002). It is possible Activin and TGF-ß actions may be mediated by different signaling intermediates (like zDYNLRB) or differently regulate the functions of some signaling intermediates (like zDYNLRB) to exert its opposing functions on ovarian follicle development. An alternative possibility is that Activin and

TGFβ may utilize a similar signaling pathway, but regulate different target genes involved in ovarian follicle development. Further studies are in need to investigate the above alternative mechanisms for the opposing effects of Activin and TGFβ ligands on ovarian follicle development.

TGFβ family signaling play important roles in early embryo development processes such as dorsal-ventral axis formation. A combination of genetic, embryological and molecular analyses in many model organisms (like Xenopus, zebrafish, and mouse) has provided much insight into the mechanisms of genetic control of this process, which is controlled by both maternal and zygotic signals (Schier and Talbot, 2005). In zebrafish, TGFβ family signals plays an important role in this process, including ventralizing signals (like BMPs), dorsalizing signals (like Nodal), and antagonists (like 163 Chordin), to counter ventralizing BMP signals (Toyama et al, 1995; Rodaway et al, 1999;

Sirotkin et al, 2000; Imai et al, 2001; Schier and Talbot. 2005; Londin et al, 2005).

Recent studies demonstrated a Smad2/3 specific phosphatase PPM1A antagonizes dorsalizing activity of Nodal signaling in vivo through dephosphorylation of Smad2/3

(Lin et al, 2006). However, the mechanism underlying the dorsal-ventral axis formation is still incompletely understood. Previous studies demonstrated that zDYNLRB is expressed from the one-cell stage through 24 hr postfertilization (Thisse and Thisse,

2004). Since the homologues of zDYNLRB in human (DYNLRB1 and 2) appear to be required for Smad2 and Smad3-dependent TGFβ signaling in mammalian cells (Ding and

Mulder, 2004; Jin et al, in revision; Chapter 3), it might be expected that their homologue zDYNLRB play a role in transducing Nodal signaling in normal dorsal-ventral axis formation, and knockdown of its expression in the embryos might disrupt the Nodal signaling, resulting in ventralized embryos. However, the preliminary results from such experiments using morphological criteria suggested that zDYNLRB might not be involved in dorsal-ventral axis formation in the zebrafish embryo, since zDYNLRB MO- injected embryos either died or underwent nonspecific development arrest at high concentrations (prelimary results in Appendix E), and display apparent normal development at low concentrations (data not shown). However, the phenotype of the dorsalization can be more accurately determined by whole mount in situ hybridization for specific molecular markers like goosecoid, although early dorsal-ventral patterning defects can be easily identified using morphological criteria by experienced scientists in the field. The results of no effects on dorsal-ventral axis formation after zDYNLRB knockdown is considered as very preliminary, since the presence of zDYNLRB protein in 164 the embryo at the one-cell stage (probably from the oocyte) may prevent such a conclusion, since such maternal zDYNLRB protein present in the embryo before the injection of zDYNLRB MOs cannot be blocked by the zDYNLRB MOs (Thisse and

Thisse, 2004). Therefore, the role of zDYNLRN in dorsal-ventral axis formation in the zebrafish embryo needs to be further determined using loss-of-function approaches in zebrafish, such as the zDYNLRB-targeted mutant zebrafish by TILLING, proposed above.

In summary, it has been demonstrated in this Chapter that the function of the zebrafish dynein light chain zDYNLRB is regulated by TGFβ in zebrafish primary ovarian follicle cells, and a potential role for zDYNLRB in TGFβ signaling in these cells.

Future studies as proposed above will investigate the physiological significance of zDYNLRB function on zebrafish ovarian follicle development and its ovarian function, like fertility. 165

Chapter 5

Overall discussion and future directions

TGFβ signaling pathways control various fundamental cellular functions (such as cell proliferation, differentiation and migration), and are essential for embryonic patterning, organogenesis, and adult tissue homeostasis. TGFβ signaling has a dual role in cancer, inhibiting tumor growth and promoting tumor progression and metastasis.

Experiments described in this thesis have shown that for the first time that DYNLRB2 is required for Smad3-dependent TGFβ signaling. Others in the lab have demonstrated a requirement of DYNLRB1 in Smad2-dependent signaling (Jin et al, in revision).

Together, these results lead us to propose a model (Fig. 25) that there is a preferential interaction between Smad3 and DYNLRB2, and between Smad2 and DYNLRB1, wherein DYNLRB2 and 1 may target Smad3 and 2 for intracellular transport by the dynein motor, respectively, thus to enhance the signaling efficiency and maintaining the signaling specificity.

There are several mechanisms to explain such preferential interactions. The first potential mechanism for such preferential interaction is that Smad3 may directly interact with DYNLRB2, and Smad2 may directly interact with DYNLRB1, and that some or all of the different amino acid residues in the protein (especially those located in the middle region) are responsible for such preferential interactions. Genetic experiments Plasma membrane TGFβ 166 P P RII RI Smad2 RB1 Smad3 RB2 ISmad2

RB2 Early endosome Early endosome Smad3 P P P P RB1 RII RI SARA II I IIRB1 Smad2 P RB2 Smad3 P P P Cytoplasm Early endosome Early endosome

P P P P II I II I I P RB2 Smad3 P P IIRB1 Smad2 P DIC DIC Dynein Dynein complex complex + + microtubule microtubule

Nucleus

Smad2 TF Smad3 Smad4 TF Smad4 167

Fig. 25. Hypothetical model. Within minutes of ligand binding, activated TGFβ receptors are internalized to early endosomes, where Smad2 is recruited by SARA to early endosomes. Once RB2 is phosphorylated by TβRII, during the same time period that Smad2 is phosphorylated by TβRI, RB2 selectively interacts with the TβR/Smad3 complex, and recruits these TGFβ signaling endosomes to the dynein motor for efficient intracellular transport through the DIC-RB2 interaction. Eventually, Smad3 is released from the dynein moter and translocates into the nucleus for transcriptional regulation of specific target genes. Similar events occur for Smad2 via its preferential interaction with RB1: within minutes of ligand binding, activated TGFβ receptors are internalized in to early endosomes, where Smad2 is recruited by SARA to early endosomes. Then RB1 is phosphorylated by TβRII, and Smad2 is phosphorylated by TβRI, respectively. RB1 then recruits the TβR/Smad2 signaling endosomes to the dynein motor through its selective interaction with Smad2. Lastly, Smad2 is released from the dynein moter and translocates into the nucleus for transcriptional regulation of specific target genes. TF: other transcription factors. 168 to test this include in vitro protein interaction experiment and in vivo protein interaction experiment. For the in vitro experiment, the proteins with appropriate tags (like GST and

Flag) may be expressed and purified in bacteria or insect cells, or prepared by in vitro transcription and translation systems, and then in vitro protein interaction experiments, such as GST pull-down assay, may be performed with the prepared proteins. Positive results would indicate direct interaction between the tested proteins to support the model.

Such direct interaction will be further confirmed when mutant proteins with the interaction domain disrupted (or key amino acids mutated in the domain) lose the ability for such interaction and/or when complementation mutations in the two proteins involved occur.

However, a major limitation of these in vitro experiments is that negative results do not necessarily mean that there is no direct interaction, due to many reasons such as that the prepared proteins may not fold properly, or may have not have proper post- translational modification (such as glycosylation, phosphorylation, prenylation, methylation and sulfation) that occur under physiological conditions in mammalian cells.

Another limitation is that proteins prepared from bacteria and insect cells (especially insect cells) may contain host proteins from the insect cells or bacteria, which may lead to false positive signal for direct interaction by work as a adaptor protein, or false negative interaction due to interfering such direct interaction, but this can be easily ruled out by running each individual purified proteins on PAGE gel and staining the gel. If additional band(s) is/are observed, then proteins will be further purified for the above experiments. 169 For in vivo experiment, FRET (fluorescence resonance energy transfer) may be performed for the detection of specific direct interactions between DYNLRB2 and

Smad3, and between DYNLRB1 and Smad2, respectively, after fusing these proteins with appropriate tags (such as GFP, YFP and the newly genetically engineered enhanced fluorescent proteins, Cerulean and Venus). FRET has gained wide use for assessing direct protein-protein interaction in vivo in living cells (Erickson et al, 2003; Giepmans et al, 2006; Kapanidis et al, 2006; Mori et al, 2004; Sturmey et al, 2006; Vogel et al, 2006;

You et al, 2006). If positive FRET signal is observed between the tested proteins when compared with control, it indicates such direct interaction. The advantage of FRET is its ability to visualize such direction interaction in living cells.

Unfortunately, FRET may require high levels of expression, thus may not represent physiological level protein-protein interaction. Therefore, the BiFC

(bimolecular fluorescence complementation) assay could be used as an alternative (Hu and Kerppola, 2003; Hu et al, 2002; Kerppola et al, 2006; Shyu et al, 2006). An advantage of the BiFC assay is that it can enable the detection of protein-protein interactions at concentrations close to their normal physiological levels, or a subpopulation of such proteins (Kerppola et al, 2006; Shyu et al, 2006). The BiFC assay is based on the reassembly of the two separated N- and C-terminal fragments of the fluorescent proteins (originally using YFP, now Venus and Cerulean) into a fluorescent complex when they are fused to proteins of interest, co-expressed and brought together by an interaction between such proteins in living cells (Hu and Kerppola, 2003; Hu et al,

2002; Shyu et al, 2006). In comparison with control, positive signals would indicate such 170 interaction between the tested proteins. Due to strong signal and direct readout, the BiFC assay has been widely accepted and used for the study of protein-protein interaction in living cells (Citovskyet al, 2006; Grinberg et al, 2004; Hu and Kerppola, 2003; Hu et al,

2002; Hynes et al, 2004; Kerppola et al, 2006; Nakahara et al, 2006; Schmidt et al, 2006;

Shyu et al, 2006). Another advantage of the BiFC assay is that there are different tags for antibody detection fused to these fragments (Flag tag for N-terminal fragment, and HA tag for C-terminal fragment), so easy confirmation of the detected protein-protein interaction can be performed by immunoprecipitation/blot and immunofluorescence staining analyses. A major limitation of the BiFC assay is that the BiFC complexes are essentially irreversible, but this would not interfere the proposed experiments here.

FRET and the BiFC assay may be performed in combination to utilize the advantages of both to test whether there is in vivo direct interaction between Smad3 and

DYNLRB2, and between Smad2 and DYNLRB1, respectively. It is essential that mutant proteins be used as negative controls in which a substitution or a small deletion is introduced into the potential interaction domain for both FRET and the BiFC assay.

The second potential mechanism for the preferential interaction between Smad3 and DYNLRB2, and between Smad2 and DYNLRB1, is that there is an indirect interaction between these proteins, and adaptor proteins bring these proteins into a protein complex. An adapter protein usually has several protein interaction domains to bind to different proteins. Assume, for simplicity, there is one adapter protein for Smad3 and DYNLRB2, and another protein for Smad2 and DYNLRB1, or one adapter with 171 different interaction domains for Smad3 and Smad2, and different interaction domains for

DYNLRN2 and DYNLRB1.

To test this potential mechanism, the adaptor protein needs to be identified and then disruption of the adaptor would lead to disruption of the above preferential interaction. First, the protein complex needs to be purified by either immunoprecipitation, or purification through a protein purification tag, or combination of both. For purification of the protein complex by immunoprecipitation, specific antibodies for the endogenous proteins are required. Smad3 and Smad2 antibodies are commercially available, and several antibodies for DYNLRN1 can potentially be used for immunoprecipitation (from preliminary results in the lab), but there is no antibody for

DYNLRB2 available yet for immunoprecipitation, which needs to be developed for such experiment). For purification through a protein purification tag, these proteins need to be genetically engineered to have a tag for purification with commercially available reagents. Protein purification tags include GST tag, His-tag, and streptavidin binding peptide purification tag. Proteins expressed with such tags can be easily purified with corresponding methods. For example, proteins with streptavidin binding peptide purification tag have a high affinity (at nanomolar level) for streptavidin resin and mild, specific elution of native proteins can be effectively eluted with biotin.

After protein complex is purified, it is prepared for PAGE electrophoresis, and each individual band on the gel is excised, followed by digestion with trypsin. The digestion products from each individual band are separated by 2D separation devices or by HPLC (High Performance Liquid Chromatography). Several such separated digestion 172 products from each individual band can be obtained individually, for example, by excising the individual spot on a 2D gel or collection of individual peaks from HPLC.

The last step is to perform mass spectrometry (MS) or tandem mass spectrometry

(MS/MS) on these individual digestion products for protein identification.

After identification of the adaptor protein(s), similar analyses on the direct interaction between the adaptor protein(s) and Smad3 (or Smad3, or DYNLRN2, or

DYNLRB1) will then be performed to confirm the role of such adaptor proteins, as proposed for testing the first potential mechanism.

The third potential mechanism for the preferential interaction between Smad3 and DYNLRB2, and between Smad2 and DYNLRB1 is that there is/are molecule(s) responsible for sorting the Smad3 and DYNLRB2 proteins located on early endosomes, and the Smad2 and DYNLRB1 proteins located on early endosomes, into distinct pools so that they are regulated by distinct pathways and drive distinct responses.

Genetic experiments to test this mechanism require the identification of the sorting molecule(s). Such molecules may be identified by screening library of mutant cells, as has been performed for TGFβ receptor (Laiho et al, 1990 and 1991) and Dab2 in libraries of chemically mutagenized cells (Hocevar and Howe, 1996; Hocevar et al,

2001). First, a library of mutant cells will be made using the piggyBac DNA transposon.

Recently, it has been demonstrated that piggyBac efficiently transposes in human and mouse cell lines, and it is the most efficient (highly flexible and active) transposon, as 173 compared to Sleeping Beauty, Tol2, and Mos1 transposon systems in four mammalian cell lines (Ding et al, 2000; Wu et al, 2006).

Advantages of transposon piggyBac to be utilized here include: 1. Transposition events distribute widely throughout the genome, with a bias for landing in transcription units, where they create loss-of-function mutations; 2. The genes disrupted by the transposon can be easily identified by PCR, eliminating the need for mapping the locus of such genes; 3. The piggyBac transposon can easily mediate the introduction of 14 kb foreign DNA in mammalian cell lines, which may include multiple genes (such as hygromycin resistance and Venus genes) to facilitate selection of stable cell clones and visible observation of such clones; 4. The inserted piggyBac transposon can be precisely excised from their original insertion sites in the genome, allowing the reversion of insertional mutations and the mutant phenotypes. This transposon system permits for the first time the efficient production of insertional mutants in the mouse, and has been used to generate 75 different genetic knockout mouse strains in three months (Ding et al, 2005;

Harris, 2005). This system now is being used to producing the first comprehensive genome-wide set of knockout mouse mutants (Normile, 2006).

To make a library of mutant cells, a two-component piggyBac transposon co- transfection system will be constructed, consisting of both a donor and a helper plasmid to detect piggyBac-mediated chromosomal integration events in cultured cells. The donor plasmid contains the piggyBac elements in which the piggyBac transposase is replaced by hygromycin resistance resistance gene, and Venus gene. The helper plasmid contains a CMV promoter-driven piggyBac transposase fragment to allow its ubiquitous 174 expression, but lacked the terminal sequences required for transposition, thus making piggyBac transposase unable to be introduced into the genome of the cells. Therefore, the transposition events from the donor plasmid will be one time, without further introducing the piggyBac transposase. The hygromycin resistance gene and Venus gene may be put under the control of SV40 promoter.

Then, co-transfection of the two plasmids will be made in the HaCaT cells, since a requirement of DYNLRB2for Smad3-dependent signaling is demonstrated in this cell line in Chapter 3, and previous reports show that distinct Smad3-depdendent and

Smad2-dependent pathways exist in this cells (Frederick et al, 2004; Kim et al, 2005;

Kretschmer et al, 2003; Levy and Hill, 2005). Hygromycin resistant stable clones will be obtained for further analyses after hygromycin selection.

Next, whether there is disruption of the preferential interaction between Smad3 and DYNLRB2, or between Smad2 and DYNLRB1, in any of the mutant stable cell clones, will be observed by automated LUMIER analyses in 96-well plates on robotics platform (Barrios-Rodiles et al, 2005), through a collaboration with Dr. Jeffrey L.

Wrana’s laboratory. Manual LUMIER analyses have been successfully used in the lab

(some results presented in Chapter 3). The automated LUMIER analyses will require transient transfection of the mutant stable clones in 96-well plates with hRL-Smad3 and

DYNLRB2-Flag, or with hRL-Smad2 and DYNLRB1-Flag, as previously described

(Barrios-Rodiles et al, 2005). Or hRL-Smad3 and DYNLRB2-Flag, or hRL-Smad2 and

DYNLRB1-Flag, may be put in the piggyBac elements of the donor plasmid, thus saving the trouble for transient transfection later. Such stably transfected mutant cells, or the 175 mutant cells after transient transfection, will be treated with TGFβ. Then automated

LUMIER analyses will be performed. Expected results are that the TGFβ induced preferential interaction between Smad3 and DYNLRB2, and between Smad2 and

DYNLRB1, will be disrupted in some mutant stable cell clones.

Lastly, the disrupted gene(s) will be identified by performing inverse PCR to recover sequence adjacent to the ends of integrated piggyBac, sequencing such sequences and Blast search in the NCBI database. The function of the identified may be further confirmed by introducing the piggyBac transposase in the helper plasmid (in order to excise the transposon from the original insertion sites in the genome) into the corresponding mutant cells by transient transfection and restoring the phenotype, and by manual immunoprecitiation/blot analyses in the original identified mutant cell clones and such restored mutant cells. The above restored-restored cells are still mutant cells, since the transposon is just moved and integrated in other sites in the genome, but now the cells are a population of heterogeneous cells with the transposon integrated in multiple sites in the genome.

No matter which potential mechanism, or a combination of any two of the above mechanisms, is actually used by the cells for the preferential interactions, the function of DYNLRB2, DYNLRB1, the identified adaptor protein(s), and/or the sorting molecule(s) needs to be further confirmed by future experiments disrupting these proteins and observing the effect on the translocation of Smad3 and Smad2, respectively, and by making knockout mouse, or even conditional knockout mouse to further dissect their roles in the Smad3-dependent and Smad2-dependent distinct pathways. 176 Disruption of the Smad3-DYNLRB2 interaction would lead to Smad3 transport and translocation defect, and at least some phenotype overlap with the Smad3 knockout mice. For example, Smad3-deficient mice exhibit reduced fertility and defective ovarian folliculogenesis, compared with wild-type mice, indicating the requirement of Smad3 in ovarian follicle development (Tomic et al, 2002 and 2004). In addition, Drosophila mutants lacking the homologous gene ofDYNLRB2 (robl deletion mutants) display a female infertility phenotype (Bowman et al, 1999). Therefore, a phenotype with defective ovarian follicle development, and/or ovarian function and female fertility might be expected for knockout mice targeting the DYNLRB2 gene. Table 1in Chapter 1 lists the knockout mouse phenotypes of various TGF signaling components. It is also possible that other phenotypes may be observed, such as defective immune function, accelerated wound healing, or colorectal cancer, as observed in Smad3 knockout mice (Ashcroft et al,

1999; Yang et al, 1999; Zhu et al, 1998), or other phenotypes not observed in Smad3- knockout mice might be observed, since DYNLRB2 might have other currently unknown function(s) independent of Smad3.

The major limitation for the data presented in this thesis is lack of data to support the interaction between the endogenous Smad3 and DYNLRB2 proteins, since over-expression of DYNLRB2 may not accurately reconstitute the functional state of the endogenous molecules with respect to its interaction with other proteins. Therefore, one important experiment in the future is to development DYNLRB2 antibodies and to identify specific antibodies for detection of such endogenous interaction to provide further evidence to support the proposed model. 177 Another major limitation for this thesis is the lack of physiological effect and significance of the findings. Thus, other important experiments in the future are to address the role of DYNLRB2 in growth inhibition of normal cells and to address its role in tumor progression and metastasis. To determine its function in vivo in the future,

DYNLRB2 knockout mice (or conditional knockout mice) should be made, as well as double knockout mice for both DYNLRB2 and Smad3 by crossing the DYNLRB2 knockout mice with Smad3 knockout mice, and phenotype analyses will then be performed on such knockout mice. Similarly, zebrafish with targeted mutagenesis in zDYNLRB may be made by TILLING (for Targeting Induced Local Lesions in

Genomes) (Amsterdam et al, 2006; Berghmans et al, 2005; Wienholds et al, 2003) to further determine the role of zDYNLRB in zebrafish ovarian follicle development and ovarian function (such as fertility).

Other important future directions to address are how the Smad3 cargo is released, whether there is any other cargo(s) for DYNLRB2 and what is the destination of such cargo(s). In the case of the mechanism for the release of the Smad3 cargo, it would be of interest to determine whether phosphorylation or dephosphorylation of DYNLRB2 play a role, since a recent report demonstrates that specific phosphorylation of TcTex-1 at its

S82 residue is responsible for the dynein complex disassembly to release the cargo rhodopsin (Yeh et al, 2006), while we have demonstrated that phosphorylation of

DYNLRB2 is important for its recruitment into the dynein complex (Chapter 3).

Other potential cargo(s) for DYNLRB2 or DYNLRB1 for intracellular transport may be TGFβ receptor TβRII, TβRI, and other downstream signaling components (like

JNK and ERK). It has been shown that lysosome-mediated degradation regulate TGFβ 178 signaling (Anders et al, 1997 and 1998; Dore et al, 1998; Zhang et al, 2004). DYNLRB family DLC may be involved in this regulation by targeting TGFβ receptors for transport by the dynein motor from early endosomes to late ensomes/lysosomes for degradation to terminate signaling. If this is true, decreased TGFβ receptor degradation and enhanced

TGFβ signaling might be expected.

In the case of other downstream signaling components (like JNK and ERK), they may be targeted by DYNLRB2 or DYNLRB1 for transport to late endosomes for optimal signaling, as has been demonstrated for p14/MP1 (Kurzbauer et al, 2004). Since it has been shown recently the structure of DYNLRB1 are very similar to that of p14/MP1, although they have very different amino acid sequences (Ilangovan et al, 2005; Liu et al,

2006c; Song et al, 2005b). It would be of interest to see whether TGFβ regulates the recruitment of ERK and/or JNK by DYNLRB1 and/or to endosomes for optimal signaling.

Therefore, a lot of work needs to be done to further determine the function of this family dynein light chains. 179

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Appendices

The following appendices represent preliminary data that would need to be confirmed or expanded, but are included here for the purpose of providing additional support for the model proposed. 210

TGFβ (-) TGFβ (+) 80000 MDCK II

incorporation 8000070000 70% MDCK 72% 95% 70% 71% 100%7000060000 94% 76% 60000 50000

thymidine 80% * 50000

H]- 40000

3 74%

[ 60%40000

of 30000 * 40%30000 20000 20000 20% 1000010000

Percentage 0% 00 11 22 33 44 55 66 77 88 Mock + - - - + - - - Scrambled siRNA - + - - - + - - dDYNLRB1 - - + - - - + - dDYNLRB2 - - - + - - - +

Appendix A. Blocking dDYNLRB2 partially impairs TGFβ-mediated DNA synthesis inhibition in MDCK cells with high passage numbers (25-30), but not in MDCK cells with low passage numbers (<15). MDCK cells (high passage numbers) or MDCK II cells (low passage numbers) were plated in 24- well plates at 1.0X104 /cm2. They were either mock-transfected, or transiently transfected with 12 pmol/well scrambled siRNA, or dDYNLRB1 siRNA, or dDYNLRB2 siRNA. 24 h after transfection, the cells were washed and incubated with SF medium for 1 h, prior to an incubation of 24 h in the presence or absence of TGFβ (10 ng/ml). [3H]-thymidine incorporation analyses were performed as previously described (Jin et al, 2005). The asterisk (*) indicates a statistically significant difference (Student’s t-test, p<0.01) in the inhibition of DNA synthesis by TGFβ between dDYNLRB1 siRNA-transfected and scrambled siRNA-transfected MDCK cells, and between dDYNLRB2 siRNA-transfected and scrambled siRNA-transfected MDCK cells.

212

MDCK II 0.6

level 0.5 rRNA 0.4 18s by expression 0.3

0.2 normalized

dDYNLRB2 0.1

Mock + ------scrambled siRNA - + - + - + - dDYNLRB2 siRNA - - + - + - + 2.5 pmol/well 10 pmol/well 30 pmol/well

Appendix C. dDYNLRB2 siRNA specifically knockdown endogenous dDYNLRB2 mRNA expression. MDCK II cells were either mock-transfected, or transiently transfected with increasing amount (2.5, 10, 30 pmol/well in 12-well plate) of scrambled control siRNA (cat# D-001206-13-05, from Dharmacon), or dDYNLRB2 stealth siRNA (5’-AAGAGGATCCAGAGCCATAAA -3’, synthesized by Dharmacon). Real-time quantitative RT-PCR analysis of hDYNLRB2 mRNA expression (forward primer: 5'- TGG TCG TAA ATG CAG AAG G-3'; reverse primer: 5'-TAC TGT GCT CTT GGC TTT CAT C-3') from MDCK cells was performed as described in “Materials and Methods.” Results depict the average and standard deviation of dDYNLRB2 mRNA levels normalized to control 18S rRNA levels. 213

MDCK II MDCK scrambled siRNA + - scrambled siRNA + - dDYNLRB1siRNA - + kDa dDYNLRB1siRNA - + kDa 14 14 DYNLRB1 DYNLRB1 6 6

Blot Ab: DYNLRB1 Blot Ab: DYNLRB1

98 98 DIC DIC 62 62

Blot Ab: DIC Blot Ab: DIC

Appendix D. dDYNLRB1 siRNA specifically knockdown endogenous dDYNLRB1 protein expression. MDCK cells and MDCK II cells were plated in 12-well plate. 24 h later, the cell were transiently transfected with the scrambled control siRNA (cat# D-001206-13-05, from Dharmacon) (lane1: 40 pmol/well), or dDYNLRB1 siRNA (5’- AAATTATGGTTGCACCAGATA-3’, synthesized by from Dharmacon) (lane 2: 40 pmol/well). 28 h after transfection, cells were harvested on ice for western blot. Top panel, blockade of endogenous DYNLRB1 was analyzed via Western blotting with rabbit DYNLRB1 anti-serum (1:500). Bottom panel, equal loading was confirmed by blotting with an anti-DIC Ab. 214

Appendix E. Nonspecific development arrest of 24 h and 48 h embryos injected with the ATG MO. Wild-type zebrafish embryos were injected with different doses zDYNLRB ATG MO (as indicated) at the one- to two-cell stage. 24 hours and 48 hours after injection, embryos were observed and photographed. Control injections were done with injection buffer. VITA

Guofeng Gao

Education:

2000-2007 Pennsylvania State University College of Medicine, Hershey, PA Ph.D. Genetics

1995-1998 Shanghai Second Medical University, Shanghai, China M.S. Molecular Endocrinology

1990-1995 Binzhou Medical College, Shandong, China B.S. Medicine

Professional Association:

American Society of Cell Biology American Association for Cancer Research

Publications:

1. Jin, Q., Gao, G. F., and Mulder, K. M. (2007) Involvement of km23 dynein light chains in TGFβ signaling. Transforming Growth factor-Beta in Cancer Therapy (Editor: Dr. Sonia Jakowlew), The Humana Press Inc., Totowa, NJ. Review. In Press.

2. Jin, Q., Gao, G. F., and Mulder, K. M. Requirement of a new dynein light chain in Smad3-dependent TGFβ signaling. The Journal of Biological Chemistry, In revision.

3. Ilangovan, U., Ding, W., Zhong, Y., Wilson, C. L., Groppe, J. C., Trbovich, J. T., Zuniga, J., Demeler, B., Tang, Q., Gao, G. F., Mulder, K. M., and Hinck, A. P. (2005) Structure and Dynamics of the Homodimeric Dynein Light Chain km23. J Mol Biol. 352,338-54.

4. Jin, Q., Ding, W., Staub C. M., Gao, G. F., Tang, Q., and Mulder, K. M. (2003) Requirement of km23 for TGFbeta-mediated growth inhibition and induction of fibronectin expression. Cell Signal. 17, 1363-72.

5. Tang, Q., Staub, C. M., Gao, G. F., Jin, Q. Y., Ding, W., Wang, Z., Aurigemma, R. E., and Mulder, K. M. (2002) A novel TGFβ receptor-interacting protein that is also a light chain of the motor protein dynein. Mol Biol Cell. 13, 4484-96