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Home , Activin and inhibin, Decorin, INHBC

Activin receptors in gonadotrope cells: new tricks for old dogs

By

Carlis Rejón G.

Department of Pharmacology and Therapeutics McGill University Montréal, Canada

June, 2012

A thesis submitted to McGill in partial fulfilment of the requirements of the degree of Doctor of philosophy

Copyright ©Carlis Rejón G., 2012

Dedication

I have had many role models during my life: Mother Teresa, a little woman with a huge spirit, who created a worldwide institution destined to assist the poorest people, regardless of their beliefs. Marie Curie an example of perseverance and dedication towards science, she was the first person to win two Nobel prizes (in Physics and Chemistry). However, my closest source of inspiration comes from a strong woman whom, with a lot of effort was able to raise four children alone. She taught me to be perseverant, honest and humble in the pursuit of my dreams. I dedicate this manuscript to her, Carmen Gonzalez, my mom.

2 Abstract Activins are members of the transforming growth factor β (TGFβ) superfamily. Though originally identified as stimulators of pituitary follicle-stimulating (FSH) synthesis and , they also play diverse biological roles, ranging from control of to regulation of immune responses. To exert their biological effects, activins and other TGFβ family members signal through a heterotetrameric complex composed of two type I (also called -like kinases or ALKs) and two type II transmembrane serine/threonine kinase receptors. Within the superfamily, ligands greatly outnumber receptors, hence multiple ligands share receptors and individual ligands can bind various receptors. For instance, activins bind to type II receptors ACVR2 and ACVR2B, leading to recruitment, phosphorylation, and activation of type I receptors (predominantly ALK4 for activins), which in turn phosphorylate downstream effectors. In my research I am particularly interested in activin stimulation of FSHβ subunit (Fshb) expression, the rate- limiting step in production of mature FSH. Here, I used the immortalized murine gonadotrope-like cell line LβT2 to study regulation of activin signaling at the receptor level in the context of Fshb transcription. Activins induce a rapid increase in Fshb mRNA levels and FSH release from pituitary gonadotrope cells, whereas the structurally-related inhibins suppress Fshb transcription by competitively antagonizing activin binding to type II receptors. Interestingly, treating pituitary cells with the translational inhibitor cycloheximide (CHX) produces an inhibin-like effect on Fshb expression. This suggests that one of the components of the activin pathway is labile and requires continued synthesis to regulate Fshb transcription. I established that ACVR2, the main type II activin receptor in gonadotropes, is rapidly turned over in a ligand-independent fashion. My data also suggest that lysosomal and proteasomal pathways are likely not involved in the initial steps of receptor degradation, and that the ectodomain of ACVR2 might be shed at the plasma membrane. Bone morphogenetic (BMP), a sub-family of TGFβ ligands that regulates Fshb transcription along with activins, can also signal through ACVR2 in addition to the BMP type II receptor (BMPR2). I investigated whether activin A could similarly signal

3 via BMPR2. I showed that activin binds BMPR2 with low affinity. Moreover, modulation of BMPR2 expression levels in LβT2 cells, by overexpression or knockdown of the receptor, affected activin-stimulated Fshb transcription. These results indicate that BMPR2 is indeed a functional activin type II receptor in gonadotropes in vitro. Activin receptor promiscuity is not limited to type II receptors. In fact, different activin subtypes (e.g., activin B) can bind at least two other type I receptors (ALK2 and ALK7) with differing affinities. Receptor heterodimerization has been described as a mechanism to generate signaling diversity. However, no heterodimers of activin type I receptors have been described to date. I studied possible combinatorial associations among various TGFβ type I receptors, using a biophysical approach. I showed that ALK4 can form potential heterodimers with ALK2 and ALK5. Overall, my thesis work described novel mechanisms whereby activin signaling can be rapidly modulated in cellulo: variations in amount of available intact ACVR2 at the plasma membrane and signaling via BMPR2, a type II receptor not previously associated with the activin pathway. Dissection of the mechanisms that govern activin function will help us to understand how alterations in these signaling systems lead to pathological conditions such as inflammatory disease, infertility, and cardiovascular disease. This knowledge may, in turn, provide insight for development of newer therapeutic strategies.

4 Résumé Les activines sont membres de la superfamille du facteur de croissance transformant β (TGFβ). Initialement identifés comme étant stimulateurs de la synthèse et sécrétion de l’hormone folliculo-stimulante (FSH) dans l’hypophyse, elles jouent également divers rôles biologiques, allant du contrôle de la différenciation cellulaire à la régulation des réponses immunitaires. Pour exercer leurs effets biologiques, les activines, tout comme d’autres membres de la super-famille TGFβ, débutent la cascade de signalisation cellulaire par interaction avec un complexe hétérotétramérique composé de deux récepteurs de type I (également appelé activine receptor-like kinases ou ALK) et deux récepteurs de type II, les quatre étant des récepteurs transmembranaires à activité sérine/thréonine kinase. Au cœur de cette superfamille, les ligands sont beaucoup plus nombreux que les récepteurs, ce qui explique qu’une multitude de ligands partagent certains récepteurs et que des ligands spécifiques peuvent se lier à des récepteurs différents. Par exemple, les activines se lient à des récepteurs de type II appelés ACVR2 et ACVR2B, conduisant au recrutement, à la phosphorylation et à l'activation de récepteurs de type I (principalement ALK4 pour les activines), ce qui à leur tour induisent la phosphorylation d’effecteurs en aval de cette cascade. Dans le cadre de mes projets de recherche, je suis particulièrement intéressée à la stimulation par les activines de la transcription de la sous-unité β du gène FSH (Fshb), l'étape limitante dans la production de la FSH bioactive. Pour ce faire, j'ai utilisé la lignée de cellules murines immortalisées modélisant les gonadotropes, appelée LβT2, pour étudier la régulation de la signalisation par les activines au niveau des récepteurs dans le contexte de la transcription du gène Fshb. Chez les cellules gonadotropes hypophysaires, les activines peuvent induire une augmentation rapide des niveaux d'ARN messager Fshb et de la sécrétion de FSH. Au contraire, les inhibines, possédant une structure apparentée, suppriment la transcription de Fshb induite par les activines, et ce par antagonisme compétitif en se liant aux récepteurs de type II. Fait intéressant, le traitement des cellules hypophysaires avec l’inhibiteur de translation cycloheximide (CHX) produit un effet similaire à l'inhibine sur l'expression du gène Fshb. Ces résultats suggèrent que l'une des composantes de la voie de signalisation des activines est labile et nécessite d’être continuellement synthétisée afin réguler la transcription de Fshb. J'ai établi que ACVR2, le principal récepteur de type II pour l’activine dans les cellules gonadotropes, est en constante et rapide regénération (turn over) d’une manière

5 indépendante du ligand. Mes données suggèrent également que les voies lysosomales et du protéasome ne sont probablement pas impliquées dans les étapes initiales de la dégradation du récepteur, et que l'ectodomaine de ACVR2 pourrait être sectionné à la membrane plasmique. Les protéines morphogénétiques osseuses (bone morphogenetic proteins ou BMP), une sous-famille des ligands TGFβ qui sont capables de réguler la transcription de Fshb comme les activines, peuvent également signaler par le biais du récepteur ACVR2, en plus du récepteur BMP de type II (BMPR2). J’ai cherché à savoir si l'activine A peut aussi communiquer via le récepteur BMPR2. J'ai démontré que l'activine se lie à BMPR2 avec faible affinité. De plus, la modulation des niveaux d'expression de BMPR2 dans les cellules LβT2, par sa surexpression ou suppression, affecte l’activité stimulatrice de l’activine sur la transcription du gène Fshb. Ces résultats indiquent que BMPR2 est en effet un récepteur de type II fonctionnel pour les ligands activines dans les cellules gonadotropes in vitro. La promiscuité des activines pour ses récepteurs ne se limite pas au type II. En effet, différents sous-types d’activines (par exemple l'activine B) peuvent se lier à pas moins de deux autres récepteurs de type I (ALK2 et ALK7) avec des affinités différentes. D’autre part, l’hétérodimérisation des récepteurs a souvent été décrite comme étant un mécanisme permettant de générer de la diversité dans la signalisation cellulaire. Cependant, aucun hétérodimère de récepteurs de l'activine de type I n’ont été rapportés à ce jour. En utilisant une approche biophysique, j'ai étudié les associations possibles entre différentes combinaisons de récepteurs TGFβ de type I. J'ai aussi démontré que ALK4 possède le potentiel de former des hétérodimères avec ALK2 et ALK5. Dans l'ensemble, cette thèse décrit de nouveaux mécanismes de signalisation des activines, lesquels peuvent être rapidement modulés in cellulo: par variation dans la quantité de récepteurs ACVR2 intacts et disponibles à la membrane plasmique ainsi que par signalisation via BMPR2, un récepteur de type II qui n’a pas auparavant été associé à la voie des activines. Disséquer les mécanismes qui régissent le fonctionnement des activines nous aidera à mieux comprendre comment des altérations dans ces systèmes de signalisation peuvent mener à des pathologies telles que l’inflammation, l'infertilité, le cancer et les maladies cardio-vasculaires. Ces connaissances pourront, à leur tour, guider la science vers le développement de nouvelles stratégies thérapeutiques.

6 Contribution of authors

Chapter 2

I designed and conducted all the experiments in this section, except: Figures 2.1A and 2.1B: provided by Catherine Ho Figures 2.1C and 2.6D: provided by Ying Wang Supplementary figure S2.1: provided by Daniel Bernard The first draft of the chapter was written by me. The final version was elaborated together with Dr. Terence Hébert and Dr. Daniel Bernard.

Chapter 3

I designed and conducted all the experiments in this section, except: SPR assays: conducted by Mark Hancock Activin A-BMPR2 binary model: provided by Thomas Thompson Figure 3.6C: Ying Wang conducted the qPCR assay. I provided the cDNA The first draft of the chapter was written by me. The final version was elaborated together with Dr. Terence Hébert and Dr. Daniel Bernard.

Chapter 4

I designed and conducted all the experiments in this section, except: Figures 4.4A and 4.4C: provided by Irina Glazkova Complementary figure S4.7: provided by Eugénie Goupil The first draft of the chapter was written by me. The final version was elaborated together with Dr. Terence Hébert and Dr. Daniel Bernard.

7 Acknowledgments

I would like to acknowledge my supervisors, Dr. Terry Hébert and Dr. Dan Bernard, for giving me the opportunity to grow as a scientist and as a person during the last 5 years. It has been an enriching experience. I wish to acknowledge their infinite help while writing this manuscript. Dan, I hope this paragraph is grammatically correct.

I also would like to thank: My advisory committee: Dr. Hans Zingg, Dr. Stéphane Laporte and Dr. Guillermina Almazán for their support and opportune guidance. Special thanks to Dr. Almazán who was my advisor since I started working at McGill. The co-authors of my manuscripts, for their assistance and contributions. The investigators, assistants and students in the Pharmacology department and elsewhere, who shared reagents, equipment and/or ideas on science and life; particularly, Dr. Paul Clarke, Dr. Guillermina Almazán, Gayane Machkalyan, Gerry Baquiran and Victor Dumas. Dr. Simon Wing, Dr. Ulla Petäjä-Repo and Dr. Nabil Seidah, for sharing their knowledge about degradation. Antoine Caron, Nathalie Ethier and Darlaine Pétrin for their patience and willingness to share their knowledge during my training process at the Lady Davis Institute and the McIntyre Building. Phan Trieu, Ying Wang and Xiang Zhou, for helping me to get the reagents I required for my experiments and always putting a smile on my face. Past and present members of Terry and Dan’s labs, for creating a nice work environment and helping me out whenever I needed it. Special thanks to my lab pals: Dr. Catherine Ho, Irina Glazkova, Darlaine Pétrin, Stella Tran, Eugénie Goupil, Dr. Paulina Wrzal, Vanessa Libasci, Beata Bak Gavrila, Shahriar Khan and Peter Zylbergold, for their support, advice and understanding. Stella Tran, for revising the abstract for this thesis and also translating it to French.

8 Dr. Fouad Sufian El-Shahabi, for helping me to develop the dendrograms showed at the introduction (except the first one!). Dr. Uri Saragovi, for his support during my first years in Canada and at the beginning of my PhD, but especially for allowing me to know four amazing women, “the feisty girls”: Jennifer Peleshok, Pooja Jain, Ljubica Ivanisevic and Teresa Lama. The McGill-CIHR Drug Discovery Training Program for the financial support, as well as the Centre for the Study of Reproduction (CSR) and McGill and Réseau Québecois en reproduction (RQR) for their travel awards.

Finally, I wish to thank my family and close friends for believing in me and encouraging me when I felt this thesis would never end.

9 Table of Contents Dedication...... 2 Abstract ...... 3 Résumé ...... 5 Contribution of authors ...... 7 Acknowledgments...... 8 Table of Contents...... 10 List of Tables...... 12 List of Figures ...... 13 Abbreviations...... 16 Chapter 1...... 20 General introduction...... 20 1. The TGFβ superfamily overview ...... 20 2. Regulation at the level of ligand...... 22 3. Regulation at the level of receptor...... 25 3.1. Post-transcriptional modifications of TGFβ receptors...... 26 3.2. Receptor compartmentalization and degradation ...... 28 3.3. Co-receptors ...... 31 3.4. Receptor oligomerization...... 32 4. Structural determinants of ligand-binding interactions ...... 34 5. Regulation at the level of effectors...... 36 5.1. Regulation of R-SMAD activity...... 38 5.2. Co-factors and co-modulators...... 39 6. Cross-talk between TGFβ signaling pathways and other receptor families ...... 40 7. Functions ...... 42 7.1. The HPG axis...... 43 7.2. Regulation of Fshb transcription by activins and inhibins...... 45 7.2.1. The activin system...... 45 7.2.2. Models used to study Fshb regulation...... 46 7.2.3. Activin signaling in gonadotropes and Fshb transcription ...... 47 8. Rationale for Thesis...... 51 Chapter 2...... 64 Rapid turnover of ACVR2 explains the inhibin-like effect of cycloheximide on Fshb transcription ...... 65

10 Chapter 3...... 106 Activin A binds and signals via bone morphogenetic protein receptor type II (BMPR2) in immortalized gonadotrope cells ...... 107 Chapter 4...... 148 Preliminary assessment of TGFβ type I receptor combinatorial interactions using Bioluminescence Resonance Energy Transfer (BRET) assays...... 149 Chapter 5: General discussion ...... 184 5.1 ACVR2 turnover in LβT2 cells...... 185 5.2 BMPR2 as activin receptor in LβT2 cells...... 187 5.3 BMPR2 ligand binding surface...... 190 5.4 Type I receptor interactions ...... 190

5.5 β2AR interaction with TGFβ receptors ...... 193 Conclusions ...... 195 References ...... 203 Appendices ...... 228 Hazardous waste managment disposal training for laboratory personnel training certificate Safe use of biological safety cabinets training certificate Workplace hazardous materials information system training certificate Principles of laboratory radiation safety (refresher) training certificate Co-authors waivers

11 List of Tables

Chapter 1 Table 1.1: Promiscuity in TGFβ family of receptors...... 62 Table 1.2: Characteristics of TGFβ and BMP’s signaling complexes assembly...... 63

Chapter 3 Table 3.1: Apparent kinetics of activin A binding to amine-coupled receptor-Fc fusions, as assessed by surface plasmon resonance (SPR) ...... 146 Supplementary Table S3.1: Primers used for BMPR2-S mutagenesis† ...... 147

Chapter 4 Table 4.1: Summary of type I receptor interactions as evaluated by BRET...... 183

12 List of Figures

Chapter 1 Figure 1.1 Dendrogram of the TGFβ ligand superfamily...... 55 Figure 1.2 Canonical TGFβ signaling ...... 56 Figure 1.3 Ribbon representation of TGFβ ligands...... 57 Figure 1.4 Type I and type II TGFβ receptors...... 58 Figure 1.5 Modes of receptor assembly in the TGFβ family...... 59 Figure 1.6 The SMAD family of proteins ...... 60 Figure 1.7 The Hypothalamic-Pituitary-Gonadal (HPG) axis...... 61

Chapter 2 Figure 2.1 Prolonged exposure to cycloheximide (CHX) blocks activin A-stimulated Fshb mRNA expression in murine gonadotropes ...... 93 Figure 2.2 CHX inhibits activin A-induced SMAD2 phosphorylation without affecting total SMAD2 protein levels in LβT2 cells ...... 94 Figure 2.3 CHX causes a time-dependent reduction in activin A binding to LβT2 cells...... 95 Figure 2.4 ACVR2 is the primary type II receptor that mediates activin signaling in LβT2 cells...... 96 Figure 2.5 ACVR2 degradation is not ligand-dependent ...... 97 Figure 2.6 Proteasome inhibition partially reverses the effect of CHX on activin pathway ...... 98 Figure 2.7 Evaluation of possible degradation pathways for ACVR2 ...... 99 Supplementary figure S2.1 CHX inhibits activin A-stimulated SMAD 2/3 phosphorylation in LβT2 cells without affecting total SMAD 2/3 levels ...... 100 Supplementary figure S2.2 CHX and MG132 effects on pSMAD2 levels in different cell lines ...... 101 Supplementary figure S2.3 Validation of activin receptor type II siRNAs ...... 102 Supplementary figure S2.4 Effect of proteasomal inhibition on ACVR2-HA status and pSMAD2 levels in LβT2 cells...... 103 Supplementary figure S2.5 Lysosome inhibition does not prevent ACVR2 degradation ....104 Supplementary figure S2.6 Effect of Brefeldin A on ACVR2-HA degradation in LβT2 cells ...... 105

13 Chapter 3 Figure 3.1 Activin A binds to BMPR2 ...... 133 Figure 3.2 A hypothetical model of activin A bound to BMPR2...... 134 Figure 3.3 BMPR2 mutants affect activin A binding ...... 135 Figure 3.4 BMPR2 ECD antagonizes activin A but not TGFβ1 signaling...... 136 Figure 3.5 Over-expression of BMPR2-S enhances activin A signaling in LβT2 cells ...... 137 Figure 3.6 BMPR2 knockdown attenuates activin A-mediated signaling in LβT2 cells .....138 Figure 3.7 Inhibitors of activin signaling counteract effects of BMPR2-S over-expression...139 Supplementary Figure S3.1 Activin A binds to ACVR2 and BMPR2 but not TGFBR2 extracellular domains ...... 140 Supplementary Figure S3.2 Expression levels of HA-tagged receptors...... 141 Supplementary Figure S3.3 Activin B signaling is also affected by BMPR2-S expression in LβT2 cells...... 142 Supplementary Figure S3.4 Over-expression of BMPR2 long isoform does not enhance activin A signaling ...... 143 Supplementary Figure S3.5 The BMPR2-L construct is expressed and functional ...... 144 Supplementary Figure S3.6 Evaluation of Bmpr2 siRNAs ...... 145

Chapter 4 Figure 4.1 Saturation curve for ALK5 homo and heterodimerization...... 172 Figure 4.2 TGFβ type I receptor homodimerization...... 173 Figure 4.3 TGFβ type I receptor heterodimerization...... 174

Figure 4.4 Co-immunoprecipitation of β2AR and ALK5 in HEK293 cells...... 175 Supplementary Figure S4.1 Testing the functionality of tagged type I receptors ...... 176 Supplementary Figure S4.2 Effect of TGFβ1 treatment on ALK5 homo and hetero- dimerization...... 177 Supplementary Figure S4.3 Relative luminescence (A) and fluorescence (B) values corresponding to the cells used in Figure 4.2...... 178 Supplementary Figure S4.4 Relative luminescence values (RLU) corresponding to the cells used in Figure 4.3...... 179 Supplementary Figure S4.5 Relative fluorescence values (RFU) corresponding to the cells used in Figure 4.3 ...... 180 Supplementary Figure S4.6 Competition of the ALK5 homodimer in the presence of different

type I receptors and β2AR ...... 181

14 Supplementary Figure S4.7 β2AR colocalize with ALK5 at the plasma membrane ...... 182

Chapter 5 Figure 5.1 BMPR2 turns over rapidly in LβT2 cells...... 199 Figure 5.2 Substitution of five residues from the ACVR2 ECD in BMPR2 confers the ability to further enhance activin A signaling...... 200 Figure 5.3 Isoproterenol treatmen t does not affect activin A-induced SMAD2 phosphorylation in LβT2 cells...... 201

Figure 5.4 Expression of β2AR followed by isoproterenol treatment reduces TGFβ1 induced CAGA-luc reporter activity...... 202

15 Abbreviations ACTB: β-actin ACVR2: Activin receptor, type II ACVR2B: Activin receptor, type IIB ADAM: A disintegrin and metalloproteinase ALK: Activin receptor-like kinase AMH: Anti-Müllerian hormone AMHR2: AMH receptor, type II ARIP: Activin receptor-interacting protein BACE: Beta-secretase 1 BMP: Bone morphogenetic protein BMPR2: BMP receptor, type II BRET: Bioluminescence resonance energy transfer CamKII: Ca2+/calmodulin-dependent protein kinase II CBP: CREB binding protein CCP: Clathrin-coated pits cDNA: Complementary deoxyribonucleic acid CGA: Chorionic α CHIP: C-terminus of Hsc70 interacting protein CREB: cAMP response element-binding DMEM: Dulbecco’s Modified Eagle Medium dNTP: Deoxynucleotide triphosphate Dpp: (the Drosophila ortholog of mammalian BMPs) DTT: Dithiothreitol EDTA: Ethylenediaminetetraacetic acid EGF: EMT: Epithelial to mesenchymal transition ER: Endoplasmic reticulum ERK: Extracellular signal-regulated kinase FBS: Fetal bovine serum FOXL2: Forkhead box L2

16 FP: F2α receptor FSH: Follicle-stimulating hormone FSHβ: Follicle-stimulating hormone, β-subunit FST: GAPDH: Glyceraldehyde 3-phosphate dehydrogenase GDF: Growth and differentiation factor GFP: Green fluorescent protein GnRH: Gonadotropin-releasing hormone GnRHR: Gonadotropin-releasing hormone receptor GPCR: G protein-coupled receptor GSK3: Glycogen synthase kinase-3 GTPase: An enzyme that can hydrolyze guanosine triphosphate (GTP) HA: Hemagglutinin HaCaT: Human cells HEK293: Human embryonic cells HPG: Hypothalamic-pituitary-gonadal IFN-γ; IL-1: Interleukin 1 IP: Immunoprecipitation I-SMAD: Inhibitory SMAD JNK: c-Jun N-terminal kinase kb: Kilo basepair LAP: Latent associated LH: Luc: Luciferase MAPK: Mitogen-activated protein kinase MG101: N-acetyl-leucinyl-leucinyl-norleucinal (or calpain inhibitor-I) MG132: N-(benzyloxycarbonyl)-leucinyl-leucinyl-leucinal MH: Mad MMP1: Matrix metalloprotease 1 mRNA: Messenger ribonucleic acid

17 MTMR4: Myotubularin-related protein 4

NH4Cl: Ammonium chloride NK-1R: Neurokinin 1 receptor PAR: Protease-activated receptor PASMC: Pulmonary artery smooth muscle cells PBS: Phosphate-buffered saline PCR: Polymerase chain reaction PCA: Protein complementation assay PDP: Pyruvate dehydrogenase phosphatase PDZ: A domain named after PSD-85, Dlg and ZO-1 proteins; mediates protein-protein interactions PI3K: Phosphatidylinositol 3-kinase PKC: Protein kinase C PLB: Passive lysis buffer PMA: Phorbol 12-myristate 13 acetate PP1c: Type 1 serine/threonine protein phosphatase PP2A: Serine/threonine protein phosphatase 2A PPM1A: Protein phosphatase 1A PTH: PTH-1R: PTH type I receptor PVDF: Polyvinyl difluoride qRT-PCR: Quantitative reverse transcription PCR RACE: Rapid amplification of cDNA ends RET: Resonance energy transfer RIPA: Radioimmunoprecipitation assay Rluc: Renilla luciferase ROCK: Rho-associated protein kinase Rpl19: 60S ribosomal protein L19 R-SMAD: Receptor SMAD SARA: SMAD anchor for receptor activation SBE: SMAD binding element

18 SDS-PAGE: Sodium dodecyl sulfate polyacrylamide gel electrophoresis SMAD: Mothers against decapentaplegic homolog Smurf: SMAD ubiquitination-related factor SnoN: Ski-related novel protein N SPC: Subtilisin-like proprotein convertase SUMO: Small ubiquitin-like modifier TACE: TNF-α converting enzyme (also known as ADAM17) TAK1: TGFβ-activated kinase 1 TAP: Tandem affinity purification TBS: Tris-buffered saline TGFβ: Transforming growth factor β TGFBR2: TGFβ receptor, type II Tiul1: TGIF-interacting ubiquitin ligase 1 TKV: Thick-veins. Dpp type I receptor in Drosophila TNF-α: alpha USP15: Ubiquitin-specific peptidase 15 WT: Wild type

19 Chapter 1 General introduction 1. The TGFβ superfamily overview

The transforming growth factor β (TGFβ) superfamily consists of a group of structurally related, secreted polypeptides characterized by the presence of two subunits stabilized through hydrophobic interactions, and in most cases by an inter-subunit disulfide bond (see also Section 2) [1, 2]. Despite their high structural similarity, members of the TGFβ family share only ∼25% amino-acid sequence identity [2]. To date, more than 30 ligands have been identified in the TGFβ family, including three isoforms of the eponymous TGFβs, 10 bone morphogenetic proteins (BMPs), activins, inhibins, 11 growth and differentiation factors (GDFs), and anti-Müllerian hormone (AMH) (Figure 1.1), [3]. The majority of these ligands function as homodimers, but some of them are also able to form heterodimers.

Some TGFβ family ligands are ubiquitously expressed (including TGFβ itself), while others have a more restricted tissue distribution (for example, /GDF8). These ligands can function in an autocrine, paracrine, or endocrine fashion to control diverse biological processes, including tissue , , neuronal growth, reproductive function, the epithelial to mesenchymal transition (EMT) and immunity [3, 4]. At the cellular level, they modulate cell growth, differentiation, migration, adhesion and in a spatio-temporally regulated manner [3]. These roles are particularly relevant during early embryogenesis, when members of TGFβ family function as morphogens determining cell fates depending on their local concentrations [5] and contributing to the differentiation of germ layers. Due to their pleiotropic functions, disturbances in TGFβ family signaling are implicated in a variety of pathologies such as cancer, skeletal malformation, fibrosis, wound healing dysfunction, autoimmune and cardiovascular disease [4, 6-8].

To exert their diverse biological functions, TGFβ family ligands signal through oligomeric complexes formed by two groups of structurally related serine-threonine

20 kinase-containing receptors, termed type I (or activin-like receptor kinases - ALK) and type II receptors, according to their relative molecular masses [9-15]. Dimeric ligands can initially bind to high affinity sites located on either type I (e.g., BMP2 or BMP4) or type II receptors (BMP7, activins and TGFβs), which is followed by recruitment of complementary low affinity binding receptors (corresponding type II or I, respectively) and subsequent formation of heteromeric complexes [10, 11, 16, 17]. The exact stoichiometry of the different receptor complexes is unknown at present. Although traditionally considered to be a heterotetramer, composed by two type II and two type I receptor dimers [18], more recent data suggest that type I/type II heterodimers may represent independent signaling units [19].

Regardless of the receptor combination, once the ternary signaling complex is formed, type II receptors trans-phosphorylate the type I receptors, which then undergo a conformational change resulting in subsequent activation of their own protein kinase domains [20-22]. The activated type I receptor transmits intracellular signals through transient association and activation of downstream effectors, of which members of the Mothers Against Decapentaplegic/SMA (SMAD) related family of proteins have been the most intensely studied [23].

The SMAD family of transcription factors contains eight members in mammals (see also Section 5). According to their function, SMADs are classified as receptor-regulated or R- SMADs (SMADs 1, 2, 3, 5 and 8), the common or co-SMAD (SMAD4) and inhibitory or I-SMADs (SMADs 6 and 7) [24]. The activated type I receptors interact with R-SMADs and phosphorylate them on a C-terminal SSXS motif. Receptor-mediated phosphorylation of R-SMADs prompts their association with SMAD4 and subsequent accumulation into the nucleus. Once in the nucleus, the SMAD complexes bind to various promoter elements whereby they positive or negatively regulate expression of different genomic targets (Figure 1.2). The minimal SMAD binding cis-element (SBE) has been defined as 5’-GTCT-3’ and its reverse-complement 5’-CAGA-3’ [25], although phosphorylated SMAD1 interacts preferentially with GC-rich sequences (such as 5’- GCCGNCGC-3’ or 5’-GGCGCC-3’) [26]. These GC rich sequences are usually in close

21 proximity to the canonical SBE, allowing SMAD4 partners to make additional contacts with the DNA [27-29]. Given that SMADs bind DNA with low affinity or, in the case of SMAD2, do not directly bind DNA at all [30, 31], they associate with sequence-specific DNA binding proteins to modulate transcriptional activity (see also Section 5.2). In addition to this canonical signaling cascade, diverse SMAD-independent pathways can be activated by TGFβ superfamily. For instance, TGFβ can induce ERK1/2 activation, as well as JNK, p38 and PIK3 signaling [32-37].

Altogether, TGFβ family members activate a seemingly straightforward linear signaling cascade, where ligands bind to two types of receptors (type I and type II) activating downstream effectors (SMADs) which regulate gene transcription. This canonical pathway, in conjunction several alternative pathways, allows TGFβ superfamily members to initiate and control a diverse array of functions. Surprisingly, in contrast to the large number of ligands present in the family and their pleiotropic roles, only a limited number of receptors (five type II and seven type I) have been described in mammals. Therefore, many strategies have evolved to generate signaling diversity. At the same time, given the relevance of TGFβ superfamily activity during embryogenesis and in the mature organism, it is not surprising several mechanisms also exist to control the duration, intensity, and specificity of TGFβ family signaling.

Over the next sections, I will present an overview of the cellular mechanisms that exist at the level of the ligands, receptors and effectors (SMADs), to modulate signaling of TGFβ superfamily members. I will emphasise regulation at the level of the receptors since this subject is the main focus of my thesis.

2. Regulation at the level of ligand

TGFβ superfamily members comprise homo- or heterodimeric ligands. Each subunit is often referred to as a “hand”, with two fingers (nine β strands arranged in two antiparallel β sheets), a “thumb” (the N-terminus), a wrist (a long α-helix perpendicular to the β strands) and a “knuckle” region (the convex side of the β sheets), held together by three

22 or four disulfide bridges, which form a motif called a “cysteine knot” (Figure 1.3) [1, 2]. In the dimeric form, the majority of the ligands (including TGFβ1, TGFβ2 and BMP7) show a symmetric extended arrangement that evokes a butterfly-like shape, where each a monomer represents a wing [38, 39]. However, certain members, such activin and TGFβ3 are flexible and can adopt alternative conformations [38, 40].

TGFβ ligands are synthesized as pre-proproteins that contain a large amino-terminal pro domain region (called latent associated peptide – LAP) and the C-terminal mature ligand. The formation of a biologically active dimeric growth factor occurs intracellularly and requires at least three steps: a) correct folding of monomers, including the generation of the cysteine knot motif through intra-subunit disulfide bonds; b) dimerization of the pro- proteins and c) proteolytic cleavage of the N-terminus pro-region by proteases of the subtilisin-like pro-protein convertase (SPC) family at an Arg-X-X-Arg recognition site [29, 41, 42]. In this sense, heterodimer formation presumably occurs at the pro-protein stage rather than in the mature proteins. Many of ligands in the family are secreted as latent forms, noncovalently associated with their pro-domains, and require further processing by different proteases such as BMP1/Tolloid to become active. [42]. Nevertheless, both nodal and BMP9 precursors can bind and activate their respective receptors [43-45] while the pro-form of BMP2 competes with mature BMP2 for receptor binding, but does not signal [46]. Therefore, in certain instances TGFβ ligands action can be regulated by the availability of the relevant activating proteases.

The activity of TGFβ ligands can be further controlled by secreted proteins that associate with their mature forms, neutralizing their effects. Among these soluble extracellular antagonists are: a) , /SOG and DAN/, which bind BMPs and are relevant for the establishment of proper BMP gradients during early embryogenesis; b) the and α2-macroglobulin that bind TGFβ; and c) follistatin (FST)- follistatin-related gene (FLRG) which bioneutralize activins, GDFs and BMPs [47, 48].

FST is a cysteine-rich single chain with no sequence similarity to TGFβ family ligands. It exists as three different isoforms generated by alternative splicing and

23 partial proteolysis. These isoforms (FST-288, FST-305 and FST-315) are named after the number of residues they contain. According to data generated using X-ray crystallography, two molecules of FST encircle a single molecule of activin, blocking its type I and type II receptor binding sites [49]. FST also inhibits BMP signaling to a lesser extent than activins, by interacting simultaneously with both BMP and a type I receptor [50].

The presence of local diffusible antagonists and ligand-activating proteases plays a key role for the establishment of ligand gradients. Since members of the TGFβ superfamily (activins and BMPs) act as morphogens during embryogenesis [51], variations in local antagonist and proteases levels can elicit different outcomes [5, 52]. There are different mechanisms that explain how the intensity of a stimulus (ligand concentration) might induce the expression of different . These mechanisms are reviewed elsewhere [29, 52, 53].

In addition to modulating ligand bioavailability, another way to generate TGFβ signaling diversity is through ligand heterodimerization. Most members of the TGFβ family of ligands have been studied as homodimers due to the relative ease of generating purified ligands. However several ligands are able to form heterodimers, including activins (see Section 7.2.1), BMP 2/7, BMP 4/7, BMP 2/6, TGFβ1/2, TGFβ2/3, BMP15/GDF 9 and inhibins [54-61]. The majority of ligand heterodimers seem to signal in a similar fashion to their corresponding parental homodimers, however certain heterodimers possess unique signaling properties. For example BMP4 and BMP7 homodimers are weaker inducers of mesoderm markers in Xenopus and osteogenic differentiation in primary cultures of rat bone marrow stromal cells, compared with the BMP4/7 heterodimer [58, 59]. In a similar fashion, BMP2/7 dimers seem to induce ventral mesoderm, while the BMP-2 homodimer is incapable of doing so [57]. Also, BMP2/6 heterodimers are more potent than its homodimeric counterparts to induce human embryonic differentiation [56]. These differences in functionality might be due to the recruitment of distinct receptor combinations or differences in affinities for the same receptor complex.

24 Therefore, coexpression of different ligand subunits can lead to more potent responses, or in some cases, to new biological effects.

In summary, ligand bioavailability is determined by the levels of soluble antagonist, and activity of ligand processing enzymes. In turn, variations in the local concentration of active ligand and ligand heterodimerization can alter the intensity of a given effect or generate a spectrum of biological responses in a temporal- or cell-specific manner.

3. Regulation at the level of receptor

As discussed above (see Section 1), TGFβ superfamily members bind to two different types of serine/threonine kinase-bearing receptors classified as type I (65-70 kDa) and type II (85-110 kDa). To date, five type II receptors (TGFBR2, BMPR2, AMHR2, ACVR2, ACVR2B) and seven type I receptors (ALK1-7) have been identified in mammals (Figure 1.4A). In addition, BMPR2, ACVR2B and TGFBR2 have different isoforms. Some TGFβ members can also bind accessory molecules at the plasma membrane called co-receptors, which can modulate access of ligands to the signaling receptors (see Section 3.3).

The type I and type II receptors share a similar gross structure, characterized by a short, highly variable, ligand-binding extracellular amino-terminal chain, a single-pass transmembrane domain, and a large cytoplasmic region comprised mainly of a serine/threonine kinase domain. The crystal structures of the extracellular domains (ECDs) of four type II receptors (TGFBR2, ACVR2, ACVR2B, BMPR2) have been determined [38, 40, 62, 63]. As expected from sequence analysis, these receptors exhibit a similar overall topology, with the presence of a three “finger” toxin fold (a commonly observed fold found in snake venom neurotoxins and cardiotoxins) composed of three different pairs of antiparallel β-strands connected by four conserved inter-sheet disulfide bonds (Figure 1.4B). The β-strands are curved so as to generate a concave surface for ligand binding. The crystal structures of ALK3 and ALK5 also show the “three-finger toxin” fold observed in the type II receptors (Figure 1.4C) [39, 64].

25

Despite their overall architectural resemblance, there is little amino acid sequence identity between type I and type II receptors, especially in the extracellular domains [2, 39, 65]. Besides sequence divergence per se, other differences distinguish type I from type II receptors. For instance, the type II receptors are protein kinase active dimers, which suggests the possibility that they might be autophosphorylated; whereas type I receptors need to be phosphorylated by type II receptors in a glycine/serine-rich domain (called the GS box) in order to be activated. Type I receptors also possess a short amino- acid sequence called the L45 loop, which mediates their interactions with certain downstream effectors (see Section 5) [1, 22, 66, 67] . On the other hand, type II receptors contain a relatively short C-terminal extension following the kinase domain absent in type I receptors. One BMPR2 isoform (BMPR2-L) possesses a distinctive long C- terminal tail with 530 amino acids after the kinase domain [9], which likely fosters interactions with other proteins [68-71]. The functional relevance of the C-terminal tail extension for other type II receptors has not been determined, but seems to be unnecessary for basic [72].

As we will see over the next sections, the activity of type I and type II receptors can be modulated by reversible translational modifications such as phosphorylation, sumoylation and ubiquitination. In addition, compartmentalization of the receptors, the presence of accessory proteins (co-receptors) and receptor oligomerization contribute to the generation of signaling diversity and provide additional control points for signaling regulation.

3.1. Post-transcriptional modifications of TGFβ receptors

Phosphorylation of type I receptors in the juxtamembrane GS domain results in a conformational change that reduces the affinity of the receptors for an inhibitory regulatory molecule called FKBP12, while favouring its interaction with downstream effectors [20, 21]. Therefore, phosphorylation within the GS box is crucial for type I receptor activation and subsequent triggering of signaling pathways. In fact, mutagenesis

26 of the penultimate residue of the GS box (either a threonine or glutamine residue) into acidic amino acids (aspartate or glutamate), thus mimicking the negative charge of a phosphorylated residue, constitutively activates the type I receptor, and allows propagation of intracellular signals in the absence of either ligand or type II receptors [15, 73, 74].

Consistent with the dynamic nature of protein phosphorylation, several phosphatases have been identified, that can inhibit receptor activity. For instance, the type 1 serine/threonine protein phosphatase 1 (PP1c) can modulate activity of Dpp type I receptor (Tkv) in Drosophila [75] and dephosphorylate ALK1, ALK3 or ALK5 in mammalian cells [76-78]. Likewise, the Ser/Thr protein phosphatase 2A (PP2A) interacts with ALK3-BMPR2 even in the absence of ligand, and attenuates BMPR2 phosphorylation [79]. Thus, the balance between receptor phosphorylation, induced by the presence of ligand, and the action of these phosphatases, will determine the extent of the biological responses and helps control spurious signaling in the absence of stimuli. TGFBR2 and ALK5 can be also phosphorylated on tyrosine residues following TGFβ stimulation, favouring the activation of mitogen-activated protein kinase (MAPK) pathways [80, 81]. However, the tyrosine phosphatases that reverse this modification have not yet been identified.

Another post-translational modification reported for ALK5, but no other type I (or type II) receptors is sumoylation (i.e. the covalent attachment of a SUMO molecule to lysisne residues). Sumoylation of ALK5 in response to TGFβ enhances recruitment and activation of SMAD3, increasing ligand responsiveness [82]. ALK5 sumoylation requires prior activation of the receptor by phosphorylation [82], therefore it is regulated by the activation state of type I and also type II receptors.

Receptor phosphorylation and sumoylation provide different ways to modulate receptor function and delineate cellular responses, by controlling the activation of both SMAD and non-SMAD signaling pathways. In addition to these two types of post-translational

27 modifications, the type I receptors can also be ubiquitinated and targeted for degradation as we will see in the next section.

3.2. Receptor compartmentalization and degradation

Receptor endocytosis provides another means to dampen cellular signaling, and to modulate cell responses in the presence of ligand. At present, two primary internalization mechanisms have been described for TGFβ receptors- clathrin-coated pit (CCP)-mediated endocytosis and internalization via non-coated vesicles (including lipid rafts and caveolae). TGFβ superfamily receptors are constitutively internalized via CCP and can be recycled to the plasma membrane through a Rab11-dependent process [83, 84]. Different motifs and residues have been identified in both type I and II receptors necessary for their internalization [85-87]. In addition, receptor internalization can be facilitated by the presence of interacting proteins. For instance, a PDZ protein called ARIP2 (activin receptor-interacting protein 2) can enhance ACVR2 endocytosis, while a co-receptor called betaglycan or TGFBR3 (see Section 3.3) can promote ALK6 and TGFBR2 internalization in a β-arrestin 2-dependent manner [88-90]. Although the majority of the ligand-unbound receptor seems to be constitutively internalized and recycled back to the plasma membrane [91], activated receptor complexes can have different fates once they are internalized.

Endocytosis of activated receptors via CCP is usually associated with an increase in SMAD phosphorylation and SMAD-dependent signaling due to enrichment, in early endocytic compartments, of scaffold proteins that stabilize SMAD binding to receptors (see Section 5) [84, 92, 93]. In this sense, inhibition of clathrin-dependent trafficking may reduce TGFβ-induced SMAD2 activation [92]. Nevertheless, internalization is not essential for TGFβ, activin or BMP signaling [87, 94], given that early receptor-effector interaction occurs at the plasma membrane [84, 92, 94]. Clathrin-dependent internalization of activated receptors can also lead to their lysosomal degradation [83, 95, 96]. It is unknown, at present, which variables regulate endosomal sorting of receptors between these destinations (signaling and recycling versus degradation), but it seems that

28 the association of receptor complexes with different proteins contributes in determining their final fate. For example, a molecule called Dapper (Dpr2) facilitates trafficking of ALK4 and ALK5 into lysosomes [97], while the viral E3 ubiquitin ligase K5, and perhaps the mammalian E3 ligase Itch, serve a similar role for BMPR2 [98].

Receptor internalization via lipid rafts/caveolae can generate different outcomes, but is normally associated with a decline in ligand signaling. Endocytosis of TGFβ receptors via caveolae can promote their interaction with complexes formed by the I-SMADs (SMAD6, SMAD7) and E3 ubiquitin ligases (such as Smurf 1 and 2 or Tiul1), resulting in receptor ubiquitination and subsequent proteasomal degradation [92, 99-102]. Opposing the action of Smurfs and other ubiquitin ligases, at least two enzymes (UCH37 and USP15) have been reported to deubiquitinate ALK5 and inhibit its degradation [103, 104]. The equilibrium between the ubiquitination states of the receptor can regulate the duration of TGFβ signaling.

In addition to mediating protein internalization, caveolin-1 can interact directly with the receptors at the plasma membrane and affect cellular responses. Caveolin-1 binds to ALK5 and inhibits TGFβ-induced SMAD2 phosphorylation [105], while, in endothelial cells, caveolin-1 associates with ALK1 to promote in the presence of TGFβ [106]. Compartmentalization of TGFβ receptors into lipid rafts has also been associated with activation of SMAD-independent signaling and the efficient formation of ligand- induced receptor complexes [84, 96, 107]. In summary, internalization of activated receptors via CCP is associated with SMAD signaling and lysosomal degradation, while internalization via lipid rafts is linked to non-canonical signaling and proteasomal degradation, although alternative signaling responses can be elicited as well.

Even though the mechanisms that control receptor dynamics are unknown, it is clear that receptor compartmentalization affects both the nature and duration of biological responses. Possibly, these effects depends on the differential expression of cellular components (I-SMADs, ubiquitin ligases, deubiquitinases, Dpp), that promote

29 degradation or signaling once receptors are internalized via lipid rafts or clathrin coated pits.

Degradation of ALK5, ALK3, ALK6 and BMPR2 can be initiated at the plasma membrane, due to the action of sheddases (membrane-bound proteases). The cleavage of activated ALK5 is mediated by TNF-α converting enzyme (TACE or ADAM17) in a PKCζ-dependent manner, leading to down-regulation of TGFβ signaling [108, 109]. Likewise, the presence of soluble forms of the BMP receptors, and a concomitant reduction in BMP2 responses, have been reported in cultures of primary human bone cells treated with phorbol 12-myristate 13 acetate (PMA), a potent inducer of sheddase activity [110]. Unfortunately, this study did not identify the enzyme(s) involved in receptor cleavage. Shedding represents an alternative way to decrease or alter receptor activity, in addition to constitutive internalization and degradation of the receptors via the lysosome or proteasome.

Although the half-life of individual TGFβ receptors has not been rigorously assessed, several studies suggest that both TGFBR2 and ALK5 are rapidly degraded. The half-life of cell surface receptors was estimated to ~1-2.5 hours for TGFBR2 and more than 12 hours for ALK5 and these turnover rates were accelerated in the presence of ligand [111- 113]. Due to constitutive internalization and rapid turnover of TGFβ receptors (see Chapter 2), cells can quickly sense variations in ligand concentration and respond accordingly. In this way, the level and extent of receptor activation correlates with ligand concentration.

Taken together, receptor internalization, compartmentalization (lipids rafts versus CCP) and degradation (via lysosome, proteasome or shedding) can either activate different pathways (SMAD-dependent or -independent cascades) or rapidly turn off signaling.

30 3.3. Co-receptors

Ligand-receptor interaction can be influenced at the membrane level by the presence of accessory molecules or co-receptors, which lack a discernible protein kinase domain, but can either potentiate or inhibit signal transduction. Betaglycan (TGFBR3) was the first coreceptor identified in ligand crosslinking experiments and probably remains the best characterized example. Betaglycan is an 853 amino-acid transmembrane proteoglycan, with a large glycosylated extracellular domain, and a short intracellular region rich in Ser and Thr residues. Betaglycan is widely expressed in adult and fetal tissues and it is naturally found as a non-covalent homodimer [66]. Initially described as TGFβ co- receptor by its ability to increase TGFβ2 binding to TGFBR2 (section 3.1) [114], it is now evident that betaglycan plays several other roles in the regulation of TGFβ family signaling. Membrane-bound betaglycan induces endocytosis of TGFBR2 and ALK6 upon interaction with β-arrestin, and in its soluble form (after shedding by MMP1), can sequester TGFβ away from signaling receptors [88, 90, 115]. Additionally, the presence of betaglycan is necessary for high affinity binding of inhibin to ACVR2 and ACVR2B, so it is recognized as an inhibin receptor [116]. It has been also reported as a BMP co- receptor, enhancing BMP binding to its type I receptors, ALK3 and ALK6 [117]. In summary, betaglycan can act as a either a positive or negative modulator of the TGFβ- mediated signaling, it is an activator of BMP function and it also promotes inhibin action while antagonizing activin-mediated signal transduction.

Another co-receptor is . Endoglin (or CD105) is a glycoprotein, structurally similar to betaglycan and shows a restricted tissue distribution, being detected mainly in hematopoietic and vascular endothelial cells [66, 118]. Endoglin enhances the interaction of TGFβ, BMP-2, BMP7 and activin with their receptor complexes, but in contrast to betaglycan, it cannot bind ligand on its own and requires association with the corresponding high affinity receptor [118-120]. In endothelial cells, membrane bound endoglin promotes angiogenesis (See Section 3.4) [121] , whereas its shed form is anti- angiogenic [122, 123]. Therefore, endoglin can either promote or prevent signaling depending on the context and its physical form (membrane-bound or soluble).

31

Co-receptors can alter cellular responses by enhancing certain ligand-receptor combinations. For instance, GPI-linked proteins from the EGF--FRL1-CFC (cryptic) family are required for nodal, GDF1, Vg1 and GDF3 signaling, as they promote association of these ligands with receptor complexes formed by ALK4/ACVR2 or ALK4/ACVR2B [124]. On the other hand, cripto prevents activin signaling by blocking its access to ALK4 [124, 125]. Likewise, RGMa and DRAGON, members of the RGM family of GPI-linked co-receptors, potentiate BMP2 and BMP4 signaling and favour its interaction with ACVR2 instead of BMPR2 (the preferred receptor in the absence of RGM proteins) [126, 127]. Some modulatory proteins can actually prevent TGFβ superfamily signaling. Among negative regulators of TGFβ signaling at the membrane level, there is the pseudo-receptor BAMBI (BMP and activin membrane-bound inhibitor), which can associate with several type I receptors (except ALK2), preventing formation of functional signaling complexes [125]; and the GPI-anchored protein CD109 that inhibits TGFβ1 signaling, presumably by modulation of ALK5 activity [128]. In summary, the presence of co-receptors may determine not only the ability of a given cell to respond to a subset of ligands but also the responses that would be elicited.

3.4. Receptor oligomerization

Since the number of ligands in the TGFβ superfamily surpasses the number of receptors identified so far (see Section 3); there is a considerable overlap in receptor usage by ligands (see Table 1.1). For instance, BMP7 and the activins bind to the same type II receptors (ACVR2 and ACVR2B, see also Chapter 3), but elicit different cellular responses mediated by recruitment of distinct type I receptors (ALK2 or ALK4, respectively, [129] . Further, ligands (alone or in the presence of co-receptors) can engage diverse receptor complexes and elicit distinct signaling outcomes depending on the particular combination of receptors present in the complex.

In addition to signaling variability generated by type I-type II receptor combinatorial interactions, assembly of type I receptor heterodimers has also been described in different

32 cell types [121, 130, 131], adding another level of organizational complexity to TGFβ systems. Both type I and type II receptor dimers can be formed in the ER in the absence of ligand and traffic as such to the plasma membrane [18]. However, recent reports suggest that type I and type II receptors exist mainly as monomers at the plasma membrane and dimerize upon ligand stimulation [132, 133]. This raises the question of whether receptor heterodimers are preformed or if their association is induced by the presence of ligands and/or modulatory proteins at the cell surface. Formation of ALK3- ALK6 heterodimers has been observed even in the absence of ligand; however BMP2 treatment leads to an increase in the number of these complexes, suggesting that a spectrum of organizational options exist [130]. Interestingly, ALK3 and ALK6, although similar at the structural level, exhibit different affinities towards BMP ligands and play dissimilar roles in a variety of biological processes [134-136]. Hence the type I heterodimer ALK3/ALK6 may represent a new functional unit, distinctive from either ALK3 or ALK6 homo-oligomers, although this hypothesis has not been explored directly. ALK3 can also dimerize with ALK2 to mediate BMP2/BMP4 signaling, this association can again take place in the absence of ligand, as determined using bioluminescence resonance energy transfer (BRET) [137]. Nevertheless, ligand- independent assembly of ALK2-ALK3 dimers under physiological conditions and their signaling properties remains to be evaluated.

Formation of type I receptor heterodimers can also be induced by the presence of accessory proteins. In endothelial cells and chondrocytes, ALK5 can either homodimerize or heterodimerize with ALK1 in the presence of endoglin [121, 138, 139]. TGFβ signaling through these distinct receptor complexes produces opposing effects. Ligand binding to TGFBR2-ALK5 in endothelial cells inhibits cellular proliferation and migration, while binding to TGFBR2-ALK5/ALK1 oligomers promotes angiogenesis [121, 138]. ALK5 can also form heterodimers with ALK2 and ALK3 in various cell types, leading to TGFβ-induced phosphorylation of SMAD1/5 in addition to SMAD2/3 signaling [131]. These authors suggested that ALK5 heterodimerization could be a widespread phenomenon and it might depend on the expression level of the receptors or the presence of specific accessory proteins [131].

33

Unfortunately less data is available regarding heterodimerization of other type I receptors or the type II receptors. To date, the only type II receptor heterodimer reported is the one formed by TGFBR2 and its splice variant TGFBR2-B [140]. In contrast to TGFBR2, TGFBR2-B seems to bind TGFβ2 with high affinity in the absence of betaglycan [141], therefore it is possible that the heterodimer may have different ligand binding properties compared to the corresponding homodimers.

Although at present the extent of type I and type II receptor combinatorial possibilities are unclear (see Chapter 4) as are the putative molecular determinants that govern such interactions, a clear conclusion from these studies is that the composition of TGFβ receptor complexes and the pathways activated vary according to cellular context. Furthermore, receptor heterogeneity generated through receptor heterodimerization and/or alternative splicing might partially explain the broad spectrum of biological responses observed in the TGFβ family, due to subtle qualitative and quantitative differences in ligand affinity or effector coupling.

4. Structural determinants of ligand-binding interactions

Both ligands and receptor ECDs from the TGFβ superfamily exhibit a number of conserved structural motifs (cystine knot and three finger toxin, respectively). However, the crystal structures of ternary complexes of TGFβ and BMP and type I-typeII receptors along with mutagenesis and in silico studies have revealed different modes of receptor assembly (Figure 1.5) [reviewed in [41, 142]]. In the case of BMPs, the concave surface of each type I receptor contacts both ligand subunits by interacting with the wrist region of one monomer and the concave β-sheet of the other [39], while the type II receptors (ACVR2, ACVR2B and BMPR2), use their concave surfaces to interact with the convex region (knuckles) of BMPs, away from the type I receptor binding surface [39, 63, 143, 144]. In the ternary structure there was no evident interaction between the ECDs of type I and type II receptors, and no major conformational changes are observed in the ligand or the receptors upon complex formation [39, 143-146]. Therefore, a sequential, rather than

34 a cooperative model for BMP receptor assembly has been proposed [142, 144, 147]. According to this model, binding of the ligand to the high affinity receptor restricts its diffusional freedom. Consequently, the ligand is concentrated in certain membrane domains increasing the probability of encountering and recruiting the low affinity receptor. This phenomenon is known as avidity (also referred as wingspread hypothesis) [142, 144, 147, 148]. Certain factors can augment ligand avidity: a) an increase in the number of receptors contacting the dimeric ligand (receptor oligomerization), which translates into an increase in the apparent affinity; and b) receptor-receptor interactions at the cell surface, probably through the transmembrane and/or cytoplasmic domains [149].

However, the avidity hypothesis is insufficient to explain the cooperative assembly of the TGFβ ternary complex, because recruitment of ALK5 (the low affinity receptor) requires the combined interaction with TGFBR2 and the ligand, but not attachment of the receptors to the membrane; that is, ALK5-ECD can bind to TGFBR2-ECD:TGFβ binary complexes in vitro [150]. In agreement with this result, the crystal structure of the ternary complex shows that an N-terminal extension of seven residues in TGFBR2 (absent in other type II receptors), forms a composite “ligand-type II receptor” interface that serves as a functional docking site for ALK5 [64]. Therefore the assembly of TGFβ tetrameric complexes is sequential and cooperative, so the interaction between TGFBR2 and TGFβ should occur before ALK5 is recruited, association of the type I receptor requires the combined ligand-type II receptor and receptor-receptor interactions. Also, in the TGFβ ternary complex, each TGFBR2 receptor uses a surface ridge formed by the base of one of its fingers to bind a cleft between the fingertips of the ligand [38], whereas binding of ALK5 to TGFβ is largely mediated by a distinctive structural motif (only present in ALK5 and ALK4), which docks with the fingers of the ligand and consequently each ALK5 monomer contacts only one ligand subunit [64, 151]. However, since ALK1/2/3 receptors have been reported to mediate TGFβ signaling, this suggests that alternative modes of TGFβ-type I receptor interaction may exist. The main differences between the ternary structures described for BMP/activin and TGFβ complexes are summarized in Table 1.2 and Figure 1.5.

35 Information about activin-receptor interactions is more limited; however the available data suggests that activin ternary complexes probably assemble in a BMP-like manner [142, 148]. Activin type II receptors bind activin on the knuckles of the fingers, rather than the fingertips [40, 144, 148] and in the current model of activin ternary complex formation, there is no direct interaction between the receptors [40, 142, 148]. However, structural data show that activin, in contrast to BMP ligands, undergoes large conformational changes when it binds to type II receptors [40, 148]. It is possible that this conformational change, induced by the interaction with ACVR2 or ACVR2B, facilitates binding of the type I receptor to the ligand [i.e., an allosteric effect of type II binding on the ligand structure allows engagement of type I [142, 148, 152]]. Resolution of the ALK4 structure will help determine whether the assembly of activin complexes is cooperative (either due to allosteric ligand-receptor interactions or direct receptor- receptor interactions) or relies on the avidity effect.

To summarize, crystallographic data and mutagenesis studies suggest that although the overall structural resemblance among ligands and receptors allow functional promiscuity in the TGFβ superfamily, minor variations in receptor/ligand architecture or their sequences are key determinants of interaction specificity and ternary complex assembly mode.

5. Regulation at the level of effectors

R-SMADs are one of the primary substrates of activated type I receptors (See Section 1) [27]. R-SMADs and SMAD4 (See Section 1) have two conserved domains, known as Mad homology domains 1 and 2 (MH1 and MH2), separated by a variable proline-rich linker region (Figure 1.6). The N-terminal MH1 domain mediates DNA binding (except for full-length SMAD2, [30, 31], nuclear translocation, and association with nuclear proteins [153, 154]. The C-terminal MH2 domain is conserved across all SMADs. It mediates SMAD oligomerization, interactions with type I receptors and association with other proteins [24, 155]. I-SMADs lack the MH1 domain, but bind to type I receptors

36 through their MH2 domains and inhibit R-SMAD signaling by various mechanisms [156, 157].

Once R-SMADs are phosphorylated by type I receptors, they undergo a conformational change that leads to their activation and association with SMAD4 [158, 159]. Receptor- SMAD interaction specificity is determined by two complementary domains, the L45 loop present in the type I receptor (Section 3) and the L3 loop located in the MH2 domain of the R-SMADs (Figure 1.6) [66, 160-162]. Based on their particular L3 loop sequences and depending on which type I receptor mediates their activation, R-SMADs are traditionally classified as TGFβ/activin/nodal-activated SMADs (SMADs 2 and 3, associated with ALK4/5/7) or BMP-regulated SMADs (SMADs 1, 5, and 8, associated with ALK1/2/3/6) [28]. However, recent findings suggest that the TGFβ system is even more complex. For instance, TGFβ can promote SMAD1/5 phosphorylation in different cell lines through ALK1/2/3 or ALK5 [121, 131, 139, 163] and BMPs can also induce SMAD2 phosphorylation under certain conditions [[164], Bernard lab, unpublished results].

Since R-SMADs must access the type I receptors to become active, the association rate between these two components will influence the extent of canonical signaling in the cell. The interaction between type I receptors and SMADs can be modulated by various proteins. On one side, there are proteins that prevent SMAD phosphorylation, such as SMAD6 and 7. The I-SMADs compete with R-SMADs for binding to type I receptors [165, 166]. SMAD7 can interact with all activated type I receptors, blocking both TGFβ/activin and BMP signaling, whereas SMAD6 preferentially inhibits BMP signaling by preventing SMAD4 recruitment [157, 166-168].

On the other hand, there are proteins that facilitate R-SMAD recognition by type I receptors. For example, SARA (SMAD anchor for receptor activation), a FYVE domain- containing scaffold protein, binds SMAD2/3 facilitating their recruitment to the receptor complex [169]. Hrs, Axin and Disabled-2 (Dab2) can also stabilize interactions between SMAD2/3 and the ALK5-TGFBR2 complex [24, 43, 47], while endofin plays an

37 analogous role to SARA in the BMP pathway [77]. Again, the balance between the opposing effects of SARA/endofin and I-SMADs affects the duration of SMAD-mediated responses. In fact, ligand stimulation induces I-SMAD expression, creating a negative feedback loop that can terminate TGFβ family signaling [168, 170, 171].

5.1. Regulation of R-SMAD activity

The functionality of activated R-SMAD is controlled by different mechanisms, such as C- terminal dephosphorylation, cytoplasmic retention or proteasomal degradation. In order to regulate gene transcription, R-SMADs must translocate to the nucleus. Phosphorylation of R-SMADs by activated receptors induces their nuclear translocation, due to exposure of a lysine-rich sequence in the MH1 domain that acts as a nuclear localization signal [154, 172]. This increase in SMAD nuclear import is accompanied by a decrease in their export rate, which leads to nuclear accumulation of activated SMADs [173]. However, as soon as receptor activity is attenuated, nuclear accumulation is lost [174], partially owing to the action of various nuclear and cytoplasmic SMAD phosphatases, such as protein phosphatase 1A (PPM1A), pyruvate dehydrogenase phosphatase (PDP) and myotubularin-related protein 4 (MTMR4) [29]. PPM1A localizes in the nucleus and constitutively dephosphorylates receptor-activated SMAD1, 2 and 3 [175, 176]. PDP1 and 2 are broadly distributed in the cell and only target SMAD1 [177, 178], while MTMR4 dephosphorylates activated R-SMADs in the early endosome, preventing translocation into the nucleus [177, 179]. Due to the opposing effects of activated receptors and constitutively active nuclear phosphatases, R-SMADs shuttle continuously in and out the nucleus during ligand stimulation, allowing the cell to constantly correlate the net amount of nuclear R-SMADs with the extent of receptor activity [29, 173, 174].

In addition to MTMR4 action, phosphorylation of SMAD linker regions by glycogen synthase kinase-3 (GSK3), MAP kinases and Ca2+/calmodulin-dependent protein kinase II (CamKII), inhibits SMAD nuclear translocation [180, 181]. Likewise, proteins such as Akt (protein kinase B, PKB) or SnoN (a transcriptional repressor), sequester SMADs in

38 the cytosol [182-184]. Therefore, nuclear accumulation of activated R-SMAD can be attenuated by inputs coming from other signaling pathways, allowing the integration of distinct cellular signals.

Finally, SMAD levels are regulated by the ubiquitin-proteasomal pathway. Several E3 ubiquitin ligases have been identified, which can target both inactive and activated R- SMADs for proteasomal degradation [181, 185-188]. Consistent with the dynamic nature of ubiquitination, a SMAD deubiquitinase has been recently reported (USP15, [189]. This enzyme, though, mainly reduces R-SMAD monoubiquitination (which prevents DNA binding) rather than polyubiquitination (which marks proteins for degradation, [189], so the search for additional SMAD deubiquitinases continues.

The stability and functionality of SMAD4 is controlled by mono- and polyubiquitination respectively. SMAD4 can be polyubiquitinated and targeted for proteasomal degradation by direct or SMAD7-mediated interactions with various E3 ligases [185, 190-192], while monoubiquitinatination in the MH2 domain (although reversible) impairs SMAD4 ability to oligomerize with activated SMAD2/3 [185, 193, 194]. Collectively, the ubiquitin- dependent degradation/inactivation of SMADs represents a rapid mechanism to decrease signaling and to selectively remove excess activated SMADs from the nucleus or inactivated SMADs from the cytosol. Together with SMAD dephosphorylation, ubiquitination contributes to dynamically regulate levels of activated SMADs, so they correspond to receptor activity. Moreover, since many of the phosphatases and ubiquitinases modulating SMAD activity and expression have different specificities, variations in the levels of these enzymes will affect distinct pathways.

5.2. Co-factors and co-modulators

Given that R-SMADs bind DNA with low affinity, they need to associate with sequence- specific DNA binding co-factors and transcription factors to modulate transcriptional activity. The list of SMAD-interacting transcription factors is vast, and includes members of the forkhead family (FOXH1, FOXO, FOXL2), zinc finger protein family (GATA3-6,

39 Schnurri), bZIP family (c-Jun, c-Fos) as well as nuclear receptors (, and glucocorticoid receptors) [see review [195]]. These interactions are both cell type- specific and context-dependent, dictating with precision, the final response to ligand stimulation.

In addition to interactions with a variety of transcription factors, SMADs can recruit diverse corepressors and coactivators to the transcriptional machinery, regulating the levels of transcriptional activity at relevant promoters. Interactors such as p300 and CBP (CREB-binding protein) increase gene transcription by relaxing chromatin structure, via their histone acetyltransferase (HAT) activity, which facilitates access of the basal transcriptional machinery [196]. Alternatively, corepressors such as the proto-oncogenes SnoN and c-Ski or the DNA binding protein TGIF, antagonize TGFβ signaling by recruiting histone deacetylases (HDACs) [197-199]. c-Ski can also compete with R- SMADs for interaction with SMAD4 and interfere with p300/CBP binding [47, 195, 197].

In general, the affinity of activated SMADs (in association with other transcription factors) for different promoter binding sites, determines the concentrations of effector necessary for a gene to be switched on or off. Therefore, the concentration of nuclear SMAD complexes, together with the expression of specific cofactors, co-repressors and co-activators with different affinities for activated SMADs and their own DNA-binding sites determine which target genes are activated or repressed.

6. Cross-talk between TGFβ signaling pathways and other receptor families

SMADs represent an important crosstalk hub between TGFβ and other signal transduction pathways. As noted above, GSK3, CamKII and MAP kinases can phosphorylate the linker region of R-SMADs and regulate their activity, while interaction with Akt affects the intracellular distribution of SMAD3 (Section 5.1). In addition, transcription of the inhibitory protein, SMAD7 can be induced by EGF, IL-1, IFN-γ and TNF-α, in a cell type-dependent manner [200]. More recently it has been shown that G

40 protein-coupled receptors (GPCRs) can trans-activate TGFβ superfamily receptors. Short treatments with serotonin (5-HT) induces ALK3 phosphorylation and SMAD 1/5/8 activation in pulmonary artery smooth muscle cells (PASMC) [201]. This effect was prevented by antagonists of the 5-HT1B receptor or inhibitors of Rho/Rho-associated- protein-kinase (ROCK) signaling [201]. Similarly, treatment of vascular smooth muscle cells (VSMC) with endothelin or thrombin leads to a transient increase in phospho-

SMAD2 levels, which was blocked by antagonists of endothelin-1 receptor (ETA/B) or the thrombin receptor (PAR-1) and by the ALK4/5/7 inhibitor, SB431542 [202-204]. These studies did not address whether changes in ALK5 phosphorylation occurred or the precise trans-activation mechanism(s) involved, but based on different results, the authors hypothesized that activation of integrins via the Rho/ROCK pathway, mediated a conformational change in TGFβ precursors (see Section 2), making more mature ligand available to interact with receptors and initiate signaling [205].

Signaling TGFβ ligands can be also regulated by direct interaction of TGFβ receptors with members of other receptor families, such as receptor tyrosine kinases and GPCRs (see Chapter 4). TrkC and the chimeric tyrosine kinase ETV6-NTRK3 bind to TGFBR2 and prevents its oligomerization with ALK5, thus inhibiting TGFβ signaling [206]. Furthermore, in the presence of parathyroid hormone (PTH), TGFBR2 phosphorylates the cytoplasmic domain of PTH type I receptor (PTH-1R), which induces their joint endocytosis [207]. Likewise, in endothelial cells, thrombin promotes endoglin and TGFBR2 internalization via PAR-1 and PKCζ [208]. This result contrasts with the effect of thrombin in VSMC, suggesting that PAR-1/TGFβ receptors crosstalk is cell specific.

On the other hand, TGFβ can affect other pathways. TGFβ treatment delays neurokinin 1 receptor (NK-1R) internalization in T cells, enhancing substance P signaling [209]; whereas in human tracheal smooth muscle cells, TGFβ1 decreases cell surface expression of β2AR [210]. Unfortunately, the mechanism(s) that govern such cell type-specific effects remains unknown. The interaction of TGFβ-elicited pathways with other signaling cascades occurs at several levels (receptors, effectors and even transcriptional targets) and are broader than what I describe here [for more complete reviews see [51, 211]]. The

41 interplay between different signaling pathways increases the complexity and diversity of TGFβ superfamily functions, but at the same time allows the cell to integrate several environmental cues to produce highly regulated responses.

7. Functions

From the previous sections, it is clear that TGFβ signaling is exquisitely regulated, from ligand availability, receptor oligomerization and compartmentalization, to combinations of different transcription factors and regulation of SMAD effectors. This tight regulation ensures the proper functioning of the TGFβ system, which is essential for embryonic development and adult tissue homeostasis [211, 212]. Indeed, gene ablation of TGFβ components (ligands, receptors, effectors) usually leads to embryonic lethality or it is associated with various skeletal malformations and fertility problems in mice knock-out models [213]. Likewise, alterations in the TGFβ system have been associated with several pathologies [for a detailed review, see [4]].

The list of biological processes regulated by TGFβ superfamily is so large that simply to enumerate them would be an ambitious task. Not only does the family comprise more than 30 members, but each of them can induce several responses in a context-dependent manner. For instance, TGFβ1 was initially described as a transforming factor due to its effects on cell growth and differentiation of normal rat kidney , but later it was shown that TGFβ can also induce inhibition of cell growth, regulate immune responses and participate in wound healing [214].

BMPs (the largest TGFβ subfamily) were discovered due to their ability to induce ectopic bone formation in muscle, although these ligands play non-osteogenic functions as well, with effects in , eye and kidney development, mesoderm induction and regulation of cardiomyocyte proliferation [215-219].

Activins also represent a good example of versatile ligands. Activins were actually isolated and linked to five different functions: stimulation of follicle stimulating hormone

42 (FSH) synthesis (for which the ligands were named), induction of erythrocyte differentiation, neuronal survival, stimulation of mesoderm development and as a non competitive inhibitor of IL-6 receptor [reviewed in [220]]. Given the broad distribution of activins (and their receptors), it is not surprising their involvement in such disparate functions and their consideration as a therapeutic target [220, 221]; however, our knowledge about regulation of activin pathways, especially at the level of receptors is still limited.

Since my work mostly focused on activin signaling, in the next few sections, I will expand on the role of activin in the context where it was first described, i.e. as a regulator of FSH . Before doing so, I need to explain the signaling network where activin is involved in regulating FSH expression, the Hypothalamic-Pituitary-Gonadal (HPG) axis.

7.1. The HPG axis

The HPG axis comprises the group of signals from the , and that regulates reproductive function in mammals. The decapeptide, gonadotropin-releasing hormone (GnRH), is secreted in a pulsatile fashion by a subset of hypothalamic neurons into the hypophyseal portal vasculature at the median eminence. The hormone then reaches the anterior pituitary (adenohypophysis), where it binds to its receptor (GnRHR), expressed in gonadotrope cells (Figure 1.7). GnRHR is a GPCR coupled to Gαq/11, therefore GnRH binding leads to activation of phospholipase C (PLC), as well as stimulation of protein kinase C (PKC), MAPK pathways and Ca2+ mobilization. The activated MAPK cascades induce expression of many genes, including the gonadotropin subunit genes (see below), whereas increased levels of Ca2+ stimulate gonadotropin secretion [222-224].

The luteinizing hormone (LH) and follicle stimulating hormone (FSH) are composed of a distinctive β subunit (that provides biological specificity) and a common α subunit (also called chorionic gonadotropin alpha or CGA) [225]. The

43 expression of the α and β subunits starts during embryogenesis (see below) and continues at a basal level until the onset of puberty, when the activation of the hypothalamic GnRH pulses enhance their expression [226]. The frequency and amplitude of GnRH pulses differentially regulates the expression of GnRHR and the gonadotropins subunits [225, 227-229]. High frequency pulses of GnRH correlate with increased Lhb (and Gnrhr) mRNA levels while slow GnRH pulses favor Fshb transcription and diminish Gnrhr expression [227-230]. The α subunit (CGA) is produced in excess and is less stringently regulated by GnRH pulses, thus transcription of β subunits is considered the rate limiting step in the synthesis of gonadotropins. GnRH pulses also control LH release, whereas the majority of FSH is sorted through the constitutive secretory pathway [231-235]. In consequence, FSH synthesis and secretion are directly coupled, so the levels of FSH in serum mainly depend on Fshb expression.

Released gonadotropins travel through the bloodstream to the gonads, where they control steroidogenesis and gametogenesis (ovarian and testicular ). In males, LH binds to Leydig cells to stimulate synthesis, while FSH regulates sperm cell maturation, although it is dispensable for male reproductive function (at least in mice) [236-238]. In females, gonadotropins act on two hormone-secreting somatic cell types called theca and granulosa cells that surround the oocyte. Theca cells respond to LH stimulation by releasing (such as and testosterone), which are converted into by adjacent granulosa cells through the actions of P450 aromatase. Aromatase expression is upregulated by FSH signaling in granulosa cells. LH also stimulates and formation of . FSH induces proliferation and differentiation and it is required for later stages of follicle growth and maturation (see next section) [239, 240]. By a negative feedback loop, the secreted sex steroids (androgens, estrogens, and progestagens), travel through the systemic circulation to the anterior pituitary and hypothalamus, inhibiting gonadotropin synthesis and secretion [241, 242].

44 In addition to GnRH input, Fshb transcription is modulated by the combined action of activin, inhibin and FST at the level of the pituitary [243-245]. Over the next sections, I will explain in more detail how activin and inhibins control Fshb transcription.

7.2. Regulation of Fshb transcription by activins and inhibins

7.2.1. The activin system

Activins and inhibins were purified from ovarian follicular fluid based on their ability to selectively modulate FSH secretion in rat primary pituitary cultures, independently of GnRH action [61, 246-248]. It is now known that activins are expressed in various tissues, including the pituitary [220, 249-251] Activins and inhibins are structurally related ligands. Inhibins are heterodimeric molecules composed of one α subunit, linked by interchain disulfide bonds to one of two distinct, but related, β subunits (βA or βB), giving rise to inhibin A (αβA) or inhibin B (αβB). On the other hand, activins are homo or heterodimers of inhibin β subunits, which form activin A (βAβA), activin B (βBβB) or activin AB (βAβB), [248, 252]. Besides βA and βB, two other β subunits (βC, βE) have been identified in mammals [253, 254] and another one in Xenopus (βD), [255]. The βC and βE subunits are expressed exclusively in the , and they do not seem to directly regulate FSH secretion [256-259].

The activin isoforms not only exhibit different distributions, but also distinct biological properties. For example, activin A only binds ALK4 (in association with ACVR2 or ACVR2B), yet activin B and the heterodimer activin AB are also able to signal through ALK7 in the presence of ACVR2 [260, 261]. In addition, activin B is less active than activin A or AB to stimulate FSH release or drive erythro-differentiation, but it is a stronger inducer of Xenopus mesoderm-differentiation [60]. Then again, the nature of the ligand might determine the potency of the response in a context-dependent manner, probably due to the recruitment of alternative receptor complexes.

45 Inhibins, on the other hand, competitively antagonize activin signaling by forming inactive complexes with the type II receptors [124]. Inhibins can bind with low affinity to ACVR2 and ACVR2B [262, 263], but in the presence of betaglycan (see Section 3.3), their affinity for the type II receptors increase, and they become potent antagonists [116, 264]. Therefore the ability of inhibin to prevent activin signaling largely depends on the expression of betaglycan in the target cell. Before explaining in more detail the activin signaling pathway in anterior pituitary, it is necessary to quickly describe the models used to study regulation of gonadotropin transcription.

7.2.2. Models used to study Fshb regulation

The study of FSH and LH transcriptional regulation was initially conducted in heterologous cell systems and primary cultures. Although a great deal of information was obtained regarding the control of gonadotropin expression, these models have limitations. Heterologous cell systems, for instance, might not recapitulate what happens in gonadotropes due to variations in the expression of cell-specific components from the GnRH, activin or steroid signaling pathways. On the other hand, primary pituitary cultures contain a heterogenous population of secretory cell types (corticotropes, somatotropes, lactotropes, gonadotropes, thyrotropes), where gonadotropes represent only 5-10% of the total population [265]. Therefore the response elicited in gonadotropes might be obscured by the paracrine effects from neighbouring cells.

To date, the only homologous models available to study the mechanism of gonadotropins transcriptional regulation are αT3-1 and LβT2 cells. These immortalized gonadotrope- like cell lines were created by targeted oncogenesis, using the 5’-promoter region of human CGA or rat Lhb to drive expression of the simian virus 40 (SV-40) T-antigen in the pituitary of transgenic mice at different developmental stages [266, 267]. The mice developed pituitary tumors from which αT3-1 and LβT2 cell lines were derived. In mice, Cga expression is activated on embryonic day 10.5 (E11.5), five days earlier than either Lhb (E16.5) and Fshb (E17.5) [268]. Therefore, LβT2 cells represent a more mature gonadotrope precursor than αT3-1. αT3-1 cells express Cga, Gnrhr and Sf1 but not Lhb

46 or Fshb. In contrast, LβT2 cells express many (if not all) characteristic markers of a fully differentiated gonadotrope including Lhb (under basal conditions) and Fshb in response to activin A [266, 267, 269-271]. However, it is important keep in mind that, being immortalized; LβT2 cells may possess distinct features from those of mature, primary gonadotropes and ideally the responses observed using this model (as well as primary cultures or heterologous systems) should be verified using in vivo models or purified gonadotrope cultures. Fortunately, the development of two different transgenic models has facilitated the isolation and enrichment of gonadotropes from mice pituitaries [272, 273], although primary cells remain difficult to expand and transfect, limiting the feasibility of certain mechanistic studies in these cells.

7.2.3. Activin signaling in gonadotropes and Fshb transcription

Numerous reports have confirmed the role of inhibins and activins on FSH regulation. For instance, intravenous or subcutaneous injection of recombinant inhibin A in rats, suppresses Fshb mRNA expression and FSH (but not LH) secretion [274, 275]. Conversely, neutralizing against inhibin α subunits or injection of purified activin A, increase circulating FSH levels [244, 276-280]. Activin A was the first activin subtype produced recombinantly in sufficient quantities for experimentation, and historically has been the most commonly used in in vitro studies [281]. There is a convincing body of evidence supporting paracrine/autocrine stimulation of FSH by activin B in the pituitary. Activin B is the predominant isoform expressed in both adult rat pituitary and in LβT2 cells [249, 282-284]. Moreover, immunoneutralization of activin B effectively inhibits FSH secretion in rat and ovine primary cultured cells and attenuates secondary FSH surge in rats [285-287]. Altogether these data suggest that activin B is the biologically relevant isoform in the pituitary for FSH regulation. Interestingly, deletion of the Inhbb gene in mice (which affects activin AB, activin B and inhibin B expression), leads to a slight increase in FSH secretion [288], suggesting that activin A from neighboring cells or other ligands might compensate for the loss of the βB subunit.

47 The availability of activin is determined by the presence of FST and other bioneutralizing proteins (Section 2) in the milieu. Indeed, levels of free activin in serum are practically undetectable, as it is mostly bound to FST [289], confirming that activins act mainly in an autocrine/paracrine fashion. In the pituitary, FST mRNA is detected mainly in folliculostellate cells (epithelioid cells that secrete ), but also in other pituitary cells (including gonadotropes) [290-293] suggesting that FST also operates locally.

In contrast to the restricted nature of activins, inhibins act mainly in an endocrine fashion. Gonadectomy results in a rapid decrease in circulating inhibin concentrations [294], which implies that testes and are the major source of circulating inhibins, although inhibin B is also found in rat gonadotropes [284, 295]. Consistent with the actions of inhibins in gonadotropes, betaglycan is expressed in rat pituitary [296] and LβT2 cells [116].

Activin initiates downstream signaling by binding to ACVR2 or ACVR2B and then recruiting the type I receptors, ALK4 or ALK7. Both activin type I and type II receptors have been detected in LβT2 cells, and primary pituitary cultures from diverse species [261, 266, 282, 297-300]. Knockout of Acvr2 in mice decreases FSH synthesis and release, without affecting LH levels [301, 302]. This result indicates that ACVR2 is the preferred type II receptor used by activins in vivo to regulate FSH transcription, and ACVR2B cannot compensate for its loss (see Chapter 3).

Likewise, knockdown of Acvr1b (ALK4) in LβT2 cells prevents activin-A induced Fshb transcription [299], while activin B and activin AB can still signal (with less potency) through ALK7 [261]. However, ACVR1C (ALK7)-deficient mice are viable and do not show fertility defects [303], suggesting that in vivo activin B signals preferentially through ALK4. Since deletion of Acvr1b gene in mice is embryonic lethal [304], the development of a gonadotrope-specific Acvr1b knockout model will be necessary to confirm this idea.

48 In LβT2 cells, activin A treatment induces SMAD2/3 phosphorylation and nuclear accumulation, followed by a rapid increase in Fshb mRNA levels [305, 306]. Activin A treatment also stimulates murine, ovine, rat and porcine Fshb promoter reporters in LβT2 cells [266, 305, 307-310]. In contrast, the human FSHB promoter is barely induced by activin A [311]. The activation of porcine and murine Fshb reporters by activin requires the presence of both SMAD 2 and SMAD3, whereas stimulation of the rat promoter involves only SMAD3 [261, 305-307, 311-313]. These results indicate that the elements involved in activin A-mediated Fshb transcriptional regulation vary among species.

Promoter analysis alone does not explain the differences observed between species. The murine and rat promoters contain a proximal consensus 8-bp SBE site, where SMAD2/3/4 complexes can bind [307, 311, 312, 314]. Instead of this element, two 4-bp minimal SBE binding sites (SB-like 1 and SB-like 2) have been identified in the proximal regions of ovine, porcine, murine, rat and even human promoters [309, 315, 316]. of the SBE sites affects activin-driven induction of murine, ovine and porcine promoters to different extents, but they are not sufficient to confer activin-sensitivity to the human promoter [309, 310, 313, 315, 316].

It is possible that the lack of response of the human FSHB promoter is due to the absence of cofactors required for full activin induction. As mentioned in Section 5.2, SMADs have low intrinsic affinity for DNA, so they need to associate with other proteins (co- factors) to increase their binding specificity and affinity for target genes. In order to play a role on Fshb transcriptional regulation, such cofactors need to be expressed in gonadotropes, exhibit a recognized binding site in the Fshb promoter and physically interact with SMAD2/3 and/or SMAD4. Among the identified SMAD-interacting partners are: PIXT1, PIXT2 and FOXL2 [317-323]. Depletion of these proteins or mutations in their DNA binding sites, impairs activin-induced Fshb promoter-reporter activity, although there are interspecies differences in their contributions to activin responsiveness [313, 315, 316, 318, 320, 322, 323].

49 Interestingly, knockdown of SMAD2/3 did not completely abolish stimulation of murine, rat and porcine Fshb promoters by activins [305, 306, 311, 313], suggesting that SMAD- independent pathways are also implicated in Fshb transcriptional regulation. One possibility is the TAK1 cascade, which might play a role in activin A-induced ovine promoter activity [324].

It is important to note that, in addition to activins and inhibins, other TGFβ ligands, the BMPs (BMP2, BMP4, BMP6 and BMP7) also modulate Fshb expression either alone or in synergy with activins, although their effects vary between species [282, 325-327].

The combined action of pulsatile hypothalamic GnRH, paracrine/autocrine activin, follistatin, BMPs, gonadal inhibin and steroids determine the changes in gonadotropin secretion during the estrous cycle in rodents [328] and the in primates [329, 330].

50 8. Rationale for Thesis

FSH action is crucial for normal reproductive function in mammals, especially in females [236, 237]. Several paracrine and endocrine factors (GnRH, sex steroids, glucocorticoids) modulates FSH synthesis; however activins are probably the most potent and selective inducers of FSH production [288, 301, 302].

In gonadotrope cells, autocrine activin signaling is tightly regulated by the endocrine feedback action of inhibins [124, 294, 331], the paracrine/autocrine effects of follistatins [290, 292, 332] and, at the intracellular level by negative modulation via SMAD7 [165, 333]. In addition, activin-stimulation of Fshb transcription requires the expression of specific transcription factors, which interact with SMAD2/3 and facilitate their binding to diverse SBE in the Fshb promoter [317-323]. Remarkably, there is much less information available regarding how activin function might be modulated by receptor availability or the particular composition of receptor complexes.

The development of the murine gonadotrope-like cell line, LβT2 has provided a suitable in vitro system to study the mechanisms controlling Fshb transcription, but also regulation of activin function, since these cells express a functional activin signaling system [300]. The objective of my thesis is to further understand how activin effects are fine-tuned in gonadotropes, in particular focusing on regulation of activin signaling at the level of the receptor. To do so, I used LβT2 cells as a model and Fshb transcription as a read-out. In Chapter 4, I also survey the potential combinatorial associations between TGFβ type I receptors using BRET in order to yield insights into the extent or limits of receptor hetero-oligomerization.

51 Figure legends

Figure 1.1. Dendrogram of the TGFβ ligand superfamily. Based on the biological function and , ligands can be classified into four sub-families as indicated: (1) BMP/GDF; (2) Activin/inhibin/nodal; (3) TGFβ and (4) Others. The background color is associated with the type of effector these ligands activate: SMAD2/3 (pink) or SMAD1/5/8 (light green). Modified from [334].

Figure 1.2. Canonical TGFβ signaling. Binding of dimeric TGFβ ligand to its type I and type II receptors (1 and 2), leads to the formation of a heteromeric complex (3) and trans-phosphorylation of type I receptor (4). In turn, the activated type I receptor phosphorylates receptor-regulated SMADs (R-SMADs) (5), allowing them to associate with SMAD4 (6) and accumulate in the nucleus (7). Once in the nucleus, SMAD complexes can interact with different co-factors (8) to modulate gene transcription (9).

Figure 1.3. Ribbon representation of TGFβ ligands. (A-B) Dimeric BMP2 molecule showing the butterfly-like architecture of TGFβ ligands, with the β-sheets emulating two fingers, the dimer interface and α-helix representing the palm and the N-terminus as the thumb. The convex side of the fingers are called knuckles, while the wrist epitope is formed by the concave side of the fingers and the palm area. (B) Is the same representation shown in (A) but viewed along the ligand’s two-fold symmetry axis. (C) Close-up of the cystine-knot. Modified from [334]. (D) Different conformations of TGFβ3. In the bound conformation, one of the ligand subunits rotates 110°, perpendicular to the intermolecular disulfide bond. Monomers are colored green and blue in each of the structures. Figures generated in Pymol [335] from PDB# 1TGK (free ligand) and 1KTZ (receptor bound). (E) Activin is a flexible ligand. Structure analysis indicates that activin can adopt an open- or closed-wing conformation, thanks to variations in the interdomain angle [40, 148]. Figures generated in Pymol [335] from PDB# 1SAY and 1NYS (open and closed conformation, respectively).

52 Figure 1.4. Type I and type II TGFβ receptors. (A) Architecture. Both groups of share a similar organization characterized by a short extracellular domain (ECD), a single-pass transmembrane domain and an intracellular domain formed mainly by a serine/threonine kinase domain. In the type I receptors, the GS box precedes the kinase domain. The dendrograms are based on the relative amino-acid sequence similarity between receptors and they were constructed using the CLC sequence viewer 6.6.2 program (algorithm: UPGMA). Only human proteins were included: ACVR2 (NP_001607.1), ACVR2B (AAH99642.1), AMHR2 (NP_065434), BMPR2 (AAH52985.1), TGFBR2 (EAW70309), ALK1 (P37023.2), ALK2 (AAH33867.1), ALK3 (AAH28383.1), ALK4 (P36896.1), ALK5 (AAH71181), ALK6 (AAH69803) and ALK7 (Q8NER5.1). (B) Ribbon representation of ACVR2 extracellular domain (ECD) showing the “three finger toxin fold”. Image generated in Pymol [335] from PDB# 1BTE. (C) Type I/type II receptors ECDs of the TGFβ superfamily adopt the same three finger toxin motif, as evidenced by an overlay of ALK5-ALK3 (left); TGFBR2-ACVR2B (middle) and ALK5- TGFBR2 (right). The three finger formed by β strands pairs are identified. Images were created in Pymol from PDB#: 2H62 (ALK3), 2L5S (ALK5), 1PLO (TGFBR2) and 2H62 (ACVR2B) [335].

Figure 1.5. Modes of receptor assembly in the TGFβ family. (A) BMP2-ALK3- ACVR2B ternary complex. The two ACVR2-ECDs (red) binds to the knuckle epitope of each ligand subunit, while the ALK3-ECD (pink) associates with the wrist region of one monomer and the palm of the other one. (B) As in (A) but rotated 90° along the X-axis, showing that there is no direct contact between the type I and type II ECD. (C) TGFβ3- ALK5-TGFBR2 ternary complex. In this case the TGFBR2-ECDs (red) interact with the finger tips of the each monomer and the ALK5 receptor ECDs (pink) bind the wrist epitope of one subunit only. (D) As in (C) but rotated 90° along the X-axis. Due to the orientation of the receptors in this case, there is direct contact between the N-termini of ALK5 and TGFBR2 (indicated by the arrows). Modified from [334]. (E) Scheme of different modes of receptor assembly for TGFβ, activin (hypothetical) and BMP.

53 Figure 1.6. The SMAD family of proteins. (A) Dendrogram representing the SMAD family of proteins. SMADs are classified according to their function as receptor-regulated (R-SMAD), common-partner (SMAD4) and inhibitory (I-SMAD). R-SMADs and SMAD4 have two MAD homology domains known as MH1 (green) and MH2 (blue). In addition, R-SMADs contain a specific phosphorylation motif (SSXS) at the C-terminus (red). Shown in pink is exon 3, observed only in SMAD2, which precludes its direct binding to DNA. R-SMADs are further classified in two subgroups according to the type of ligand that triggers their activation. In contrast to R-SMADs and SMAD4, I-SMADs have no discernible MH1 domains. The dendrogram is based on the relative amino-acid sequence similarity between SMADs and it was constructed using the CLC sequence viewer 6.6.2 program (algorithm: UPGMA). Only human proteins were included: SMAD: SMAD1 (AAB06852.1); SMAD2 (AAC39657.1); SMAD3 (AAL68976); SMAD4 (BAB40977.1); SMAD5 (AAB66353.1); SMAD6 (AAC82331.1); SMAD7 (AAL68977.1) and SMAD8 (SSI43241). (B) R-SMAD domains and some of their functions. (C) Determinants of receptor-R-SMAD specificity. The conserved MH1 and MH2 domains of R-SMADs form globular structures with surface protrusions and pockets. R-SMADs bind to DNA (purple) via a β hairpin present in the MH1 domain. The specificity of R-SMAD interactions with type I receptors is mediated by the L3 loop (and secondarily the α-helix 1 – αH-1, purple) present in the MH2 domain of the R- SMADs and the L45 loop in the type I receptors (red circle). The sequence of α-helix-2 (αH-2) specifies interactions with DNA-binding cofactors. Modified from [336].

Figure 1.7. The Hypothalamic-Pituitary-Gonadal (HPG) axis. The interactions between signals from the hypothalamus, and gonads (ovaries in females and testes in males) involved in the regulation of reproductive function in mammals, represents the HPG axis. The interactions are depicted as stimulatory (solid line, red) or inhibitory (dashed line, blue).

54 Figure 1.1

55 Figure 1.2

56 Figure 1.3

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57 Figure 1.4

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58 Figure 1.5

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59 Figure 1.6

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C

60 Figure 1.7

61 Table 1.1: Promiscuity in TGFβ family of receptors Type I receptor Type II receptor Ligand SMAD TGFBR2 TGFβ (with endoglin) ACVR2(ActRIIA) BMP-9-10 ALK1 (TSR1) ACVR2B SMAD 1, 5, 8 Activin A (ActRIIB) BMPR2 BMP-9-10 TGFBR2 TGFβ Nodal ACVR2 BMP-6-7 ALK2 Nodal SMAD 1, 5 (ActRIA/ACVR1A) ACVR2B Lefty BMP-7 BMPR2 BMP-6-7 AMHR2 AMH TGFBR2 TGFβ BMP-2-4-6-7-10 ACVR2 GDF-5-6-7 SMAD 1, 5, 8 ALK3 (BMPR1A) ACVR2B BMP-6-7 SMAD2 (?) BMP-2-4-6-7-10 BMPR2 GDF-6-7 AMHR2 AMH BMP-2-4-6-7-10 ACVR2 GDF-5-6-7 BMP-7 ACVR2B SMAD 1, 5, 8 ALK6 (BMPR1B) GDF-5 SMAD2 (?) BMP-2-4-6-7-10-15 BMPR2 GDF-6-7 AMHR2 AMH Activin A-B-A/B ACVR2 BMP-3 GDF-11 ALK4 Activin A-B-A/B SMAD 2, 3 (ActRIB/ ACVR1B) Nodal (with cripto) ACVR2B GDF-1-3 (with cripto) GDF-8-11 TGFBR2 TGFβ SMAD 1,2, 3,5 ALK5 (TβRI) BMPR2 GDF-9 SMAD 2, 3 ACVR2B GDF-8/11 Nodal GDF-11 ALK7 (ACVR1C) ACVR2B SMAD 2, 3 Activin B Activin A/B ACVR2 - ACVR2B Inhibin A/B - BMPR2 Based on information provided in [1, 121, 131, 163, 195, 337-339].

62 Table 1.2: Characteristics of TGFβ and BMP’s signaling complexes assembly

TGFβ complex BMP complex

Type II receptor ligand Finger tips of each monomer Receptors bind ligand interface “knuckles”

Concave surface of the ligand Disulfide-linked wrist and Type I receptor ligand fingers. Each receptor contacts concave β-sheet. Each receptor interface only one ligand subunit. contacts both ligand subunits

Type I receptor niche is No direct interaction between Receptor-receptor composed by a TGFBR2- type I and type II receptor ECDs interaction TGFβ interface.

Ligand lacks the recognition Pre-helix loop in the ligand is Ligand motif that motif in pre-helix loop to bind relevant for type I receptor interacts with type I type I receptor. TGFBR1 binding. receptor rotated 45º degrees.

Both receptors bind with the Both receptor types bind with Receptor ligand binding knuckles (convex surface) the concave surface of the surface curved fingers.

Sequential and cooperative. Formation of ternary complex is Assembly mode driven by avidity effect.

Ligand drives assembly of Functional tetrameric complexes functional tetrameric complex. might form in absence of ligand, Ligand relevance on presumably due to interactions receptor complex between the cytoplasmic or assembly transmembrane domains of the receptors.

Based on information provided in [64, 142, 143, 146]

63 Chapter 2 Rapid turnover of ACVR2 explains the inhibin-like effect of cycloheximide on Fshb transcription

Early studies performed in rat primary pituitary cultures showed that cycloheximide (CHX), a potent protein synthesis inhibitor, had inhibin-like effects, suppressing Fshb but not Lhb transcription [340-342]. The specific action of CHX on FSH but not LH suggested that translational inhibition might impair signaling by activins, selective regulators of FSH synthesis [61, 248]. I hypothesized that one or more components in the activin pathway is (are) rapidly turned over and therefore must be constantly replenished in order for activin to induce Fshb transcription. In this Chapter, I tried to identify this labile element(s).

64 Rapid turnover of ACVR2 explains the inhibin-like effect of cycloheximide on Fshb transcription

Carlis Rejon*, Catherine Ho, Ying Wang, Terence E. Hébert* and Daniel J. Bernard*

Department of Pharmacology and Therapeutics, McIntyre Medical Sciences Building, McGill University, 3655 Promenade Sir William Osler, Montréal, Québec, Canada H3G 1Y6

*Address correspondence to: Carlis Rejon, Daniel J. Bernard or Terence E. Hébert email: mailto:[email protected], telephone: 514-398-8803, fax: 514-398-6690 email: [email protected], telephone: 514-398-1398, fax: 514-398-6690 email: [email protected], telephone: 514-398-2525, fax: 514-398-6690

65 Abstract

Follicle stimulating hormone (FSH) and luteinizing hormone (LH) are pituitary gonadotropins that play essential roles in regulation of vertebrate reproduction. Activins and inhibins have opposing actions on FSH secretion, by either inducing or suppressing transcription of the FSH β subunit (Fshb), the rate limiting step in dimeric FSH biosynthesis. Similar to the effects of inhibins, treatment of rat pituitary primary cultures with the translational inhibitor cycloheximide (CHX) selectively suppresses FSH secretion. Using the model murine gonadotrope-like cell line, LβT2, we tested the hypothesis that a component of the activin pathway is highly labile in gonadotropes and that its rapid loss following CHX treatment leads to a decline in activin-stimulated Fshb transcription. Treatment of cells with CHX for 6 hours, but not 1 hour, completely blocked activin A-stimulated Fshb transcription. Pre-treatment of LβT2 cells with CHX for as few as 2-3 hours inhibited activin A-stimulated SMAD2/3 phosphorylation without altering total SMAD2/3 protein levels. These data indicated that CHX affects activin signaling upstream of the SMAD proteins, most likely at the receptor level. CHX rapidly and time-dependently reduced activin A binding to LβT2 cells as revealed by affinity labeling/crosslinking. RNA interference and promoter-reporter analyses showed that ACVR2 is the preferred high affinity type II receptor for activin A in LβT2 cells. Taken together, these results suggest that ACVR2 is rapidly turned over in gonadotrope cells. Indeed, overexpressed ACVR2 had a half life of ~2 h in LβT2 cells and its turnover was ligand-independent. Degradation of ACVR2 was not significantly altered by either lysosomal or proteasomal inhibitors, although inhibition of the proteasome caused accumulation of an immature form of the receptor. Preliminary data indicate that ACVR2 might be shed from the plasma membrane, but this requires further investigation. Collectively, our data suggest that CHX has inhibin-like effects in gonadotrope cells by impairing activin signaling via its type II receptor, ACVR2. Whereas inhibins competitively antagonize activin binding to the type II receptor, CHX prevents the de novo synthesis of ACVR2, which is rapidly turned over in gonadotropes.

66 INTRODUCTION

Follicle-stimulating hormone (FSH) and luteinizing hormone (LH), products of gonadotrope cells of the anterior pituitary gland, regulate gonadal development and function. Both are heterodimeric glycoproteins comprised of a common α subunit (chorionic gonadotropin α or CGA) non-covalently linked with hormone-specific β subunits (FSHβ and LHβ). The β subunits confer biological specificity to each ligand and their synthesis represents the rate-limiting step in production of active hormone [266, 343].

FSH and LH production and release are regulated by pulsatile secretion of hypothalamic gonadotropin-releasing hormone (GnRH) as well gonadal sex steroid feedback. In addition, FSH but not LH secretion, is regulated by members of the transforming growth factor β superfamily, specifically the activins and inhibins. Activins stimulate whereas inhibins suppress FSH synthesis and secretion [61, 246-248, 344].

Activins signal via binding to heteromeric complexes of type II (ACVR2 or ACVR2B) and type I (ACVR1B, or activin receptor-like kinase receptor 4, ALK4) serine/threonine kinase receptors, although some activin isoforms can also signal through ALK7 [260, 261]. Activins bind initially to type II receptors, which then trans-phosphorylate and activate type I receptors. In turn, activated ALK4 phosphorylates the downstream effector molecules SMAD2 and SMAD3. Phosphorylated SMAD2/3 interact with the canonical co-factor SMAD4, and accumulate in the nucleus where they modulate target together with cell specific DNA binding factors, such as AP1, FOXH1/FAST, Milk/Mixer and FOXL2 [305, 313, 323, 345-348]. Activins regulate FSH synthesis by driving transcription of the hormone β subunit (Fshb) mRNA [305, 307, 311, 312, 315, 343, 349] and distinct SMAD binding elements have been identified in the Fshb promoters of different species [307, 309, 311, 312, 314-316]. In addition, activins might also regulate Fshb transcription through SMAD-independent pathways [324, 350, 351].

67 In contrast, inhibins suppress Fshb mRNA levels by competing with activins for binding to type I and type II receptors [116, 352]. In the presence of the type III TGFβ receptor, also known as betaglycan, inhibins can recruit activin type II receptors (ACVR2 and ACVR2B) into non-signaling complexes, sequestering them from activin and type I receptors [116, 264, 353]. Besides preventing activin signaling, inhibins might also modulate gonadotrope function by altering the levels of GnRH receptors [245]; however, the exact mechanism and physiological relevance are unclear given that inhibins do not impact LH release [354, 355].

Initial studies designed to elucidate a mechanism of action for inhibins showed that cycloheximide (CHX), a potent protein synthesis inhibitor, mimicked inhibins’ effects in rat primary pituitary cultures [340-342]. That is, CHX markedly reduced basal FSH secretion, while LH biosynthesis and release were unaffected [341]. CHX similarly reduced Fshb mRNA levels, though its effects were reversed a few hours after inhibitor withdrawal [340, 342].

Based on these observations, the selective effect of CHX on FSH might be explained by antagonism of activin signaling in gonadotropes and/or rapid turnover of a factor involved in the stabilization of Fshb mRNA [340]. Herein, we evaluated the former possibility and examined which component(s) of the activin pathway is (are) rapidly degraded following CHX treatment using murine primary pituitary cultures and an immortalized murine gonadotrope-like cell line (LβT2). LβT2 cells possess many characteristics of gonadotropes [270] and, although endogenous Fshb mRNA expression is low in these cells under basal conditions, it is strongly induced by exogenous activin treatment [266, 269]. Our data suggest that, in the presence of CHX, ACVR2 levels quickly decrease, resulting in a rapid loss of autocrine/paracrine activin signaling and an associated decline in Fshb transcription.

68 MATERIALS AND METHODS

Reagents

Human recombinant activin A was purchased from R&D Systems (Minneapolis, MN, USA) or acquired from Dr. Tom Thompson (University of Cincinnati, OH, USA), who also generously provided follistatin 288. Enhanced chemiluminiscence (ECL) Plus reagent and Na-125I (100 mCi/mL) were purchased from Perkin Elmer (Woodbridge, ON, Canada). 1X Dulbecco’s modified phosphate-buffered saline (D-PBS) and Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 4.5g/l glucose, L-glutamine (with or without sodium pyruvate) were from Wisent (St. Bruno, Québec, Canada). Fetal bovine serum, media 199 (M199), Trizol reagent, SYBRgreen Supermix for qPCR, anti- V5 (R960-25), Lipofectamine/Plus and Lipofectamine 2000 were purchased from Invitrogen (Burlington, Ontario,Canada). Cycloheximide (CHX), n-acetyl tyrosine, urea, leupeptin, bovine serum albumin (BSA), chloramine T, potassium iodine, trichloroacetic acid (TCA), mouse monoclonal anti-HA (#H9658), EZview Red anti-HA affinity gel (E6779), EZview Red anti-myc affinity gel (E6654), phenylmethylsulfonylfluoride (PMSF), ammonium chloride, chloroquine and SB431542 were from Sigma-Aldrich Corp. (St. Louis, MO, USA). MG132 and proteasome inhibitor II were purchased from Calbiochem (La Jolla, CA, USA). Bis-sulfosuccinimidyl-suberate (BS3) was from Pierce-Thermo Scientific (Rockford, IL, USA) and G-25 Sephadex PD- 10 columns from GE Healthcare (Piscataway, NJ, USA). Protease inhibitor tablets (Complete Mini) were purchased from Roche Applied Science (Laval, Québec, Canada). D-luciferin potassium salt was obtained from BD Pharmingen (Mississauga, Ontario, Canada) and the GM6001 inhibitor from Enzo Life Sciences (CA-Brockville, ON, Canada). Microcon Ultracel-YM-10 centrifugation filter units were from Millipore (Billerica, MA, USA). The agarose A/G beads (sc-2003) were from Santa Cruz (Santa Cruz, CA, USA). Oligonucleotides were synthesized by IDT (Coralville, IA, USA). The anti-SMAD2 (#51-1300) antibody was from Zymed Laboratories (San Francisco, CA, USA). Anti-phospho-SMAD2 (#3101S) antibody was from Technology (Danvers, MA, USA). The anti-GAPDH (#AM4300) was from Ambion (Austin, TX,

69 USA); and anti-SMAD2/3 (#07-408) was purchased from Upstate Biotech (Waltham, MA, USA). The phospho-SMAD3 rabbit polyclonal antibody was a generous gift of Dr. Michael Reiss (Cancer Institute of New Jersey, New Brunswick, NJ, USA). Horseradish peroxidase (HRP)-conjugated secondary antibodies were purchased from Bio-Rad (Hercules, CA, USA). Short-interfering (si) RNAs were acquired from Dharmacon (Lafayette, CO, USA): Control (cat. # D-001210-05), Acvr2 siRNA #1 (cat. # D-040676- 01) Acvr2 siRNA #2 (cat. # D-040676-02), Acvr2b siRNA #1 (cat. # D-040629-01) and Acvr2b siRNA #2 (cat. # D-040629-02).

Constructs

The rat ACVR1B (ALK4)-HA in pcDNA3.0 expression vector was generously provided by Dr. T. Woodruff (Northwestern University, Chicago, IL, USA). To generate the C- terminal HA tagged version of the type II receptors, the ALK4 cDNA sequence was removed using HindIII and ClaI and replaced with the Acvr2 or Acvr2b cDNA sequences obtained by PCR amplification of the rat Flag-ACVR2 and Flag-ACVR2B expression constructs (also provided by Dr. T. Woodruff). We used the following primer sets (IDT, Coralville, IA, USA): Acvr2-forward 5’- GATAAGCTTGCCACCATGGG- AGCTGCTGCAAAG and reverse 5’- GATGATATCGATTAGACTAGATTCTTTGG- GAGG; Acvr2b forward 5’- GATAAGCTTGCCACCATGACGGCGCCCTGG and reverse 5’- GATGATATCGATGCTGGACTCTTTAGGGAGC. The ACVR2-HA sequence was subcloned in the pRK5-V5 vector (kindly provided by Dr. Nabil Seidah, Institut de Recherches Cliniques de Montréal, Montréal, Québec, Canada) to produce the double tagged receptor, with a V5 sequence towards the N-terminus and a HA tag at the C-terminus. ACVR2 was also subcloned into pcDNA4/myc-His vector to create a myc- tagged version of the receptor. The -1990/+1 mFshb-luc and -3740/+24 mId3-luc reporter constructs were described previously [305, 356]. The pRK5-HA-ubiquitin plasmid was a gift from Dr. Ted Dawson (Johns Hopkins University, School of Medicine, Baltimore, Maryland, USA).

70 Cell culture

The immortalized murine gonadotrope LβT2 cell line was provided by Dr. Pamela Mellon (University of California, San Diego, CA, USA), and was maintained in 10% FBS/DMEM-high glucose with sodium pyruvate. HaCaT cells were a generous gift of Dr. Anie Philip (McGill University, Montréal, Québec, Canada) and they were grown in 10% FBS/DMEM-high glucose with sodium pyruvate. Human embryonic kidney (HEK293) cells were maintained in 10% FBS/DMEM-high glucose without sodium pyruvate. HEK 293 cells stably expressing a HA tagged version of the prostaglandin F2α receptor (HA-FP) were obtained from Dr. Stéphane Laporte (MUHC Research Center) [357] and cultured in 10% FBS/DMEM supplemented with 1 μg/ml of puromycin (InVivoGen, San Diego, CA).

Primary cultures

Male C57BL6/j mice were sacrificed at 6 weeks of age in accordance with both institutional and federal guidelines. Pituitaries were extracted and primary cultures established as described [356]. 105 cells were seeded per well in 96-well plates. Cells were cultured in 10% FBS/M199 medium for 2 days and washed once with 1X PBS before treatment with CHX (0.1 or 5 μg/mL), FST-288 (2, 20 or 200 ng/mL), or SB431542 (1 μM) in 2% FBS/M199 for 24 hours. Treatments were performed in triplicate.

Reverse transcription and PCR assays

Total RNA was extracted with Trizol reagent from primary cultures or LβT2 cells seeded (2x106 cells/well) and treated in 6 well dishes, according to manufacturer`s instructions. Reverse transcription was performed on 1-2 μg of total RNA. Quantitative PCR (qPCR) assays were conducted using SYBRgreen Supermix (Invitrogen) with a Corbett Rotorgene 6000 qPCR instrument (Corbett Life Science, Kirkland, Québec, Canada). Expression of target genes was normalized relative to the gene for ribosomal protein L19

71 (Rpl19) and presented relative to the “no ligand” control condition. Sequences of qPCR primers were as follows: Fshb forward, 5’-GTGCGGGCTACTGCTACACT; Fshb reverse, 5’-CAGGCAATCTTACGGTCT; Rpl19 forward, 5’- CGGGAATCCAAGAAGATTGA; Rpl19 reverse, 5’-TTCAGCTTGTG-GATGTGCTC. In the case of RT-PCR analysis the following primer sets were used: Fshb.for, 5’- ATGAAGTTGATCCAGCTTTG; Fshb.rev, 5’-CATTTCACTGAAGGA-GCAGT; Rpl19.for, 5’-GGACAGAGTCTTGATGATCTC; Rpl19.rev, 5’-CTGAAG- GTCAAAGGGAATGTG (as previously described; [305].

Determination of follicle-stimulating hormone levels

FSH and LH levels in 20 μl of spent media from primary pituitary cultures were measured in singlet by multiplex assay at the University of Virginia Ligand Assay and Analysis Core. The sensitivity of the assay is 2.4 ng/ml for FSH and 0.24 ng/ml for LH

Activin A iodination

Activin A was iodinated with Na125I using the chloramine T method adapted from [358, 359]. Briefly, 2 μg of ligand were diluted in 10 μl of reaction buffer (0.05M Tris, 0.15M NaCl, pH 7.4) and mixed with 1 mCi of Na125I (∼17 Ci/mg) in the presence of chloramine T (20 μg/ml). After 4.5 min, the reaction was stopped by adding 20 μl 50 mM N-acetyl tyrosine, 200 μl 60 mM potassium iodide and 200 μl 0.6 g/ml urea. The labelled ligand was separated from free isotope using a PD-10 column equilibrated with 4 mM HCl, 75 mM NaCl, 0.1% BSA. Percent incorporation and protein specific activity were determined by TCA precipitation. The specific activity of the iodinated activin A was estimated to be around 25-40 μCi/μg.

Ligand binding assays

Confluent LβT2 cells plated in 6-well dishes were pre-treated for 0, 0.5, 1, 2, 3 or 5 hours with CHX (5 μg/ml) in 10% FBS-DMEM. Cells were then rinsed once and incubated 30

72 min at 37°C in Krebs-Ringer-Hepes (KRH) buffer pH 7.4 (128 mM NaCl, 5 mM KCl, 5 mM MgSO4, 1.3 mM CaCl2-2H2O, 50 mM HEPES) supplemented with 0.5% BSA. Then cells were incubated for 3.5 hours with 50 pM 125I-activin A at 4°C in KRH buffer supplemented with 0.5% BSA. The unbound ligand fraction was removed following three washes with KRH and the bound ligand was cross-linked with 0.25 mM BS3 in KRH buffer (30 min incubation at 4°C). The crosslinking reaction was stopped by rinsing the cells twice with 85 mM Tris-HCl pH 8; 30 mM NaCl. Finally, cells were lysed with RIPA buffer (1% NP-40, 1% deoxycholate, 0.1% SDS, 0.15M NaCl, 0.01 M sodium phosphate pH 6.8, 2 mM EDTA, 50 mM sodium fluoride and Complete Mini Protease Inhibitor Cocktail tablets), samples clarified by centrifugation (13,000 x g, 15 min at 4°C) and quantified using a γ counter (Packard Cobra II Auto Gamma, Perkin Elmer).

Gene reporter assays

LβT2 cells seeded in 48-well plates (at approximately 80% confluency) were transfected with -1990/+1 murine Fshb-luc (225 ng/well) and the indicated siRNAs (final concentration 5 nM) using Lipofectamine 2000 according to manufacturer’s instructions. For the gene reporter assays shown in Figure 4, siRNAs #2 were used. Twenty four hours post transfection, cells were washed once with 1X D-PBS and treated with either 1 nM (25 ng/mL) activin A in serum-free media (SFM) or with SFM alone for 24 hours. Cells were washed once with 1X D-PBS and lysed in 50 μl of passive lysis buffer (Promega). Luciferase activity was measured on an Orion II microplate luminometer (Berthold Detection Systems, Oak Ridge, TN, USA) using 20 μl of lysate and 100 μl of assay buffer (30 mM tricine, 2 mM ATP, 15 mM MgSO4, 10 mM DTT, 1 mM D-luciferin, 1 mM CoA). Treatments were performed in triplicate and experiments repeated a minimum of three times.

Immunoprecipitation (IP)

LβT2 cells plated on 6 wells and transfected with 3 μg of V5-ACVR2-HA, were placed in 1 ml of serum free media and treated with the inhibitors GM6001 (25 μM) or MG101

73 (10 μM) in the presence or absence of CHX for 5 hours. The conditioned media was concentrated around ten times with the Microcon Ultracel- YM-10 centrifugation filter units. An aliquot (10 μl) of the concentrated media was reserved (total lysate), while the rest was used for anti-V5 immunoprecipitation using protein A/G agarose beads (Santa Cruz Biotechnology, Inc), following the manufacturer instructions. To pre-clear the media we used 1 μg of control mouse IgG (Millipore) and the immunprecipitation was performed with 1-2 μl of an anti-V5 antibody (Invitrogen). The proteins were eluted by adding 40 μl of 1X Laemmli loading buffer to the beads and heating the samples 5 min at 95°C. The eluted proteins, as well as the total lysates were analyzed by immunoblot.

Immunoprecipitation (IP) under denaturing conditions

In order to prevent binding of ubiquitinated proteins to ACVR2, the immunoprecipitation was conducted under denaturing conditions. LβT2 cells plated in a 6-well format were transfected with 3 μg of ACVR2-myc and 0.5 μg of HA-ubiquitin using Lipofectamine 2000. The next day, cells were treated with MG132 or DMSO for 5 hours, then rinsed once with PBS and lysed with 100 μl of hot (90°C) denaturing buffer (50 mM Tris-HCl, pH 7.4, 1% SDS, 5 mM DTT). The lysates were collected and boiled for 5 min. To reduce the viscosity, samples were diluted with 200 μl of lysis buffer (50 mM Tris-HCl pH 7.4, 250 mM NaCl, 5 mM EDTA, 0.35% NP-40, 2 μg/mL leupeptin, 1 mM PMSF, 10 mM NaF, 1 mM β-glycerolphosphate, 1 mM Na3VO4) and sonicated for 10 sec. After removing insoluble material by centrifugation (15 min – 16,000 x g – 4°C), a fraction of the total lysate was kept at -20°C while the rest (∼250 μl) were further diluted with 700 μl of lysis buffer and used for IP assays using EZ view Red anti-myc beads (Sigma) following manufacturer instructions. In brief, 900 μl of protein lysate was incubated overnight with 50 μl of equilibrated beads on a rotating platform at 4°C. After three washes with lysis buffer, the beads were incubated with 50 μl of c-myc peptide solution (30 min at 4°C) on a rotating platform to elute gel-bound proteins. The eluted proteins, as well as the total lysates were analyzed by immunoblot.

74 Immunoblotting

In expression experiments, LβT2 cells were transfected with 3 μg of ACVR2-HA or pCDNA3.1 expression vectors using Lipofectamine 2000 in 1 mL of complete media. Cells were treated 24 hours post-transfection with CHX (5 μg/mL) in the presence or absence of different inhibitors (MG132: 10 μM; leupeptin: 50 μM, NH4Cl: 20 mM; chloroquine: 100 μM, FST: 300 ng/mL) for 5 hours. Where indicated, activin A (25 ng/mL) was also added to the media. For the time course, LβT2 cells were treated only with CHX (5 μg/mL) for different periods of time. After treatment, cells were washed once with 1X PBS and lysed with RIPA buffer. In some cases, lysates were sonicated 10 seconds at 9 W (Misonix Sonicator 3000; Mandel, Guelph, Ontario, Canada) to reduce sample viscosity. The soluble fraction was obtained by centrifugation at 13,000 x g for 15 min at 4°C. Protein lysates were subjected to SDS-PAGE and transferred to nitrocellulose filters (BioRad). Blots were probed with the indicated antibodies using standard techniques [305]. Densitometry analysis was performed using Image J (National Institutes of Health, USA).

For SMAD2 experiments, HEK293, HaCaT and LβT2 cells were seeded in 6-well plates. Once they reached 80-90% confluency, cells were placed in serum free media and treated with activin A (12.5-25 ng/mL) in the presence or absence of inhibitors (CHX: 5 μg/ml; MG132: 10 μM) as indicated in the legend figures. To validate the siRNAs used, HEK293 cells were plated in 6-well plates (1.5x105 cell/well) and transfected 2 days later with 1 μg of ACVR2-HA, 200 ng of ACVR2B-HA expression vectors along with 5 nM of the indicated short interfering RNAs (siRNAs) using Lipofectamine/Plus reagent according to manufacturer’s instructions. Then the samples were processed as indicated above.

Statistical analysis

Data presented are from either representative or pooled experiments (as indicated). Luciferase reporter data are presented as fold change from the control condition (set to

75 one) in each experiment. Differences between means were compared using one or two- way ANOVA, followed by post-hoc tests (Dunnett or Bonferroni) where appropriate (using Prism, GraphPad, La Jolla, CA, USA). In some cases, data were log-transformed prior to analyses to control for unequal variances. Significance was assessed relative to p < 0.05.

RESULTS

CHX blocks activin-stimulated Fshb mRNA expression in primary and immortalized murine gonadotropes

Prolonged exposure of rat primary pituitary cell cultures to CHX reduces FSH biosynthesis and secretion, without affecting LH levels [340-342]. Similarly, we observed inhibition of FSH, but not LH, secretion (Fig. 2.1A) and reduction of Fshb mRNA expression (Fig. 2.1B) in murine pituitary primary culture cells treated for 24 hours with CHX. Since autocrine/paracrine activins are the primary regulators of FSH biosynthesis in rat primary pituitary cultures [283, 285, 287], the selective effects of CHX on FSH versus LH suggested antagonism of activin signaling in gonadotropes. In fact, treatment of pituitary murine cultures with either the activin type I receptor inhibitor SB431542 [204] or the soluble activin bioneutralizing protein follistatin 288 [273] suppressed Fshb mRNA expression to a similar extent than CHX (Fig. 2.1B). Thus, CHX possesses properties phenotypically similar to an activin signaling antagonist in murine pituitary cultures.

To examine more directly the effects of CHX on activin signaling, we employed the immortalized murine gonadotrope-like cell line, LβT2 [266, 269]. We treated LβT2 cells with 5 μg/ml CHX for different periods of time prior to 1 hour treatment with activin A and measured levels of Fshb mRNA by RT-PCR. Activin A stimulated a robust increase in Fshb mRNA after 1 hour in control cells, but this effect was blocked in cells pre- treated with CHX for 3 or 6 hours (Fig. 2.1C). These results support the hypothesis that CHX treatment impairs activin signaling in gonadotrope cells.

76

CHX inhibits activin A-induced SMAD2/3 phosphorylation in LβT2 cells

Activin induction of Fshb transcription is SMAD2/3-dependent in LβT2 cells [305]. We therefore asked whether CHX impacts activin-regulated SMAD phosphorylation. We pre- treated LβT2 cells for different period of times with CHX prior to a 1 h treatment with 1 nM (25 ng/ml) of activin A. CHX had no effect on total SMAD2 levels within the treatment period, whereas activin A-stimulated SMAD2 phosphorylation was time- dependently attenuated (Fig. 2.2A and B). Similar results were observed with activin A- stimulated SMAD3 phosphorylation (Supplementary Fig. S2.1). To determine if CHX effect on activin-stimulated SMAD phosphorylation extended beyond LβT2 cells, we exposed HaCaT (immortalized human widely used in TGFβ studies) and HEK293 (transformed human embryonic kidney, highly responsive to activin) cells to CHX and followed by 1 h activin A (Supplementary Figure. S2.2). As LβT2, these cell lines also express endogenous ACVR2 and ALK4 [260, 360]. HaCaT cells showed a reduction in activin-stimulated phospho-SMAD2 levels after CHX treatment, whereas in HEK293 cells there was a significant reduction in basal but not activin-induced SMAD2 phosphorylation, indicating that the effects of CHX are cell-type specific. These results also suggest that CHX affects activin signaling upstream of the SMADs, perhaps at the receptor level.

CHX causes a time-dependent reduction in activin A binding to LβT2 cells

We next examined whether acute CHX treatment affects activin receptor levels in LβT2 cells. Attempts to assess endogenous receptor protein levels by western blot or pulse- chase experiments were unsuccessful due to the inadequacy of available activin receptor antibodies and/or because of low protein expression levels in these cells (data not shown). We therefore determined cell-surface receptor expression by affinity labelling with iodinated activin A. Cross-linking followed by SDS-PAGE revealed two activin-labelled complexes, corresponding to the type I and type II receptors (Fig. 2.3A). Pre-treatment with CHX for 0-5 h prior to addition of 125I-activin A lead to a decrease in ligand binding,

77 as assessed by γ-counting, and this reduction was statistically significantly after 5 hours (Fig. 2.3B). The time course of binding diminution paralleled the time course of loss of activin A-stimulated SMAD2 phosphorylation (Fig. 2.2).

Given that ACVR2 and ACVR2B are the high affinity (pM) binding sites for activin A and LβT2 cells express both receptors [266, 282], likely the CHX-induced decrease in binding is due to reduced de novo synthesis of the type II receptors. To determine the relative roles of ACVR2 and ACVR2B in activin A signaling, we transfected LβT2 cells with a murine -1990/+1 Fshb-luc reporter construct and small interfering RNAs (siRNA) that specifically target each type II receptor (Fig. S2.3A). We observed a significant decrease in both basal (LβT2 cells constitutively secrete activin B) and activin A-induced reporter activity when siRNA for Acvr2, but not Acvr2b, was transfected (Fig. 2.4A and Fig. S2.3B). The fold induction (when corrected for the decrease in basal activity) was also lower, but this did not achieve statistical significance (Fig. 2.4B). Knocking down of both receptors did not further reduce the reporter activity when comparing with cells transfected with Acvr2 siRNA alone (Fig. 2.4A and Fig. S2.3B).

Due to the low transfection efficiency of LβT2 cells [305], the functionality and specificity of the siRNAs used in these experiments were assessed by transfecting HEK293 with HA-tagged versions of rat-ACVR2 and rat-ACVR2B along with targeting siRNAs. As shown in supplementary Fig. S2.3A, the siRNAs specifically impaired the expression of their corresponding receptors, although Acvr2b siRNA #1 was not able to knock-down the expression of the overexpressed receptor due to a base mismatch between the murine and the rat sequence. The efficacy of the siRNAs was also tested on another reporter assay (supplementary Fig. S2.3C), where BMP2 drives the expression of an Id3 promoter-reporter (-3740/+24 mId3-luc)[356] upon binding to activin type II receptors or BMPR2. Again, knocking down Acvr2, but not Acvr2b, decreased reporter activity in response to BMP2.

Taken together, these data suggest that ACVR2 is the primary type II receptor in LβT2 cells mediating activin A induction of Fshb transcription. Furthermore, ACVR2 seems to

78 be highly labile in these cells, accounting for CHX-mediated inhibition of activin A binding and signaling. Since we were unable to determine the half-life of the endogenous ACVR2 receptor in LβT2 cells because the lack of suitable antibodies, we transiently transfected LβT2 cells with the C-terminally HA tagged rat-ACVR2 and evaluated its disappearance following CHX treatment. As observed in the activin binding assays, there was a rapid and time-dependent loss in ACVR2-HA levels following translation inhibition (Fig. 2.4C), and this reduction was significant after only 2 hours CHX treatment (Fig. 2.4D). These results confirmed that ACVR2 is rapidly degraded in LβT2 cells.

ACVR2 degradation is ligand-independent

Taking advantage of the fact that the overexpressed ACVR2-HA seems to mimic the behavior of endogenous ACVR2, we used this construct to further examine how the receptor might be turned over in LβT2 cells. Members of the TGFβ superfamily of receptors are constitutively internalized via clathrin-coated pits even in the absence of ligand [83, 84]. In the case of TGFβ receptors (TGFBR2 or TGFBR1/ALK5), degradation is further enhanced in the presence of ligand (TGFβ1); therefore, we evaluated if the rate of ACVR2 degradation was also ligand-dependent [111, 112]. Neither addition of exogenous activin A (Fig. 2.5A-B) nor sequestration of endogenous activin B with FST-288 (Fig. 2.5C) affected levels of ACVR2-HA compared with control conditions, where no ligand or inhibitor was added. Thus, in contrast to TGFBR2, ACVR2 stability is not affected by the presence of ligand, at least not under the conditions examined here.

Inhibition of the proteasome partially prevents ACVR2 degradation and rescues CHX antagonism of activin A-induced SMAD2 phosphorylation and Fshb expression

Although endocytosed membrane proteins are typically degraded via lysosomes, some cell surface receptors can also be degraded by the proteasome [361, 362]. Both pathways

79 have been implicated in the degradation of TGFβ superfamily receptors [92, 363, 364]. To determine the mechanism(s) responsible for ACVR2 turnover, we transiently transfected LβT2 cells with ACVR2-HA and evaluated its relative expression following 5 h of CHX treatment in the presence or absence of lysosomal (leupeptin, ammonium chloride, chloroquine, E64) or proteasomal (MG132, epoxomycin, PI-II, PI-IV) inhibitors.

Inhibition of the lysosomal pathway with leupeptin, ammonium chloride (NH4Cl), chloroquine or the irreversible inhibiotor E64, did not affect levels of ACVR2 under any of the conditions examined (Figs. 2.6A-B, Fig. 2.7B and supplementary figure S2.5A-B). In order to test the efficacy of the lysosomal inhibitors, we used HEK293 cells stable expressing a HA-tagged version of the prostaglandin F2α receptor (FP-HA), which we have previously seen is degraded via the lysosome [365]. As expected, treatment of FP-

HA expressing cells for 5 hours with NH4Cl or leupeptin resulted in accumulation of the receptor, although the effect was less noticeable once the CHX was present

(supplementary Fig. S2.5C). These data indicate that NH4Cl or leupeptin can effectively block the lysosome, but still cannot slow-down ACVR2 turnover.

Likewise, proteasomal inhibition with MG132 did not prevent ACVR2 degradation in the presence of CHX (Fig. 2.6A, compare lanes 4 vs. 12 and 2.6B). However, acute exposure of LβT2 cells to MG132 alone lead to an enrichment of ACVR2, especially a ∼80kDa form of the receptor, while another slower migrating form (∼100 kDa) was marginally affected (Fig. 2.6A compare lanes 3 vs. 11 and 2.6B). Similar results were obtained with another proteasome inhibitor, epoxomycin (supplementary Fig. S2.4A). Deglycosylation assays with Endo-H and PNGaseF enzymes showed that the smaller band (80 kDa) was Endo H-sensitive; therefore representing an immature (ER-resident) form of the receptor. On the other hand, the 100 kDa band was Endo H-resistant and PNGaseF-sensitive, presumably representing the mature form of ACVR2 (data not shown). Treatment of transiently transfected LβT2 cells with brefeldin A (an agent that prevents transport of proteins out of the ER) confirmed that the 80 kDa band corresponded to the ER-resident receptor (supplementary Fig. S2.6, compare lanes 5 vs. 6). Overall, these data suggest

80 that the immature ACVR2 (at least when over-expressed) is degraded via the proteasome in LβT2 cells, while the mature form of the receptor is turned over through an alternative route.

Since proteasome targeting requires polyubiquitination (usually K48-linked chains) of the protein [366, 367], we determined the ubiquitination state of ACVR2 in basal conditions and after proteasomal inhibition. In this case we transfected LβT2 cells with ACVR2- myc and HA-ubiquitin. After treating the cells with MG132, we immunoprecipitated the receptor and examined the presence of an HA signal. As shown in supplementary Fig. S2.4B, ACVR2 was ubiquitinated and the levels of the ubiquitinated receptor were slightly increased in the presence of MG132, suggesting that proteasomal inhibition prevents degradation of the ubiquitinated protein.

Inhibition of the proteasome with MG132 prevented the CHX effects at the level of SMAD2 phosphorylation and Fshb transcription (Figs. 2.6C-D). We pre-treated LβT2 cells for 5 hours with CHX in the presence or absence of MG132 or leupeptin and then added 25 ng/ml activin A for 1 hour. Again, CHX drastically reduced activin A- stimulated pSMAD2 levels without affecting total SMAD2 levels. The proteasome inhibitors MG132, PI-II and, to a lesser extent, PI-IV (Fig. 2.6C, lanes 6 and 12; supplementary Fig. S2.4C) rescued basal and activin A-induced pSMAD2 levels; whereas leupeptin had no effect (Fig. 2.6C, lanes 5 and 11). We also evaluated the effect of MG132 on activin A-induced Fshb expression. We incubated LβT2 cells with CHX alone or with MG132 for 5 hours, followed by 2 h treatment with activin A (25 ng/ml) in the continued presence of the inhibitors. Then, RNA was isolated, and Fshb mRNA measured by qPCR (Figure 2.6D). As expected, activin A increased Fshb expression, and this induction was blocked by CHX treatment. Importantly, MG132 partially rescued the activin A response in the presence of CHX. Given that MG132 did not prevent turnover of the mature ACVR2, we consider that the effects of the proteasome inhibitors on activin cascade, might be directly at the levels of the SMADs, which are degraded by the proteasome (Fig. 2.6C, compare lanes 1 versus 3 and 7 versus 9) [92, 188, 368].

81

Is ACVR2 shed at the plasma membrane?

To further characterize the pathways involved in ACVR2 degradation, we treated ACVR2-HA expressing LβT2 cells, with inhibitors of calpain (MG101) or aminopeptidases (bestatin), and we also tested conditions that favours (serum withdrawal) [369] or prevent (treatment with 3MA inhibitor) [370] macroautophagy. Again, none of the inhibitors/conditions was able to completely prevent ACVR2 degradation once protein synthesis was inhibited by CHX (Fig. 2.7A-B, supplementary Fig. S2.4A). MG101 had a similar effect to the proteasome inhibitors in the absence of CHX, favouring the accumulation of an immature form of the receptor. We also noticed the appearance of a smaller molecular weight band (~55 KDa) in cells treated with MG101 (Fig. 2.7B, lane 5). This band also appeared in cells treated with MG132 or epoxomycin (supplementary Fig. 2.4C, lanes 3-6), and it was almost undetectable (at least not at short exposure times) in cells treated with other inhibitors.

The ∼55KDa band could represent a degradation product of the mature receptor or an unglycosylated form of full-length ACVR2. Some members of the TGFβ receptor superfamily, such as ALK3, ALK6, ALK5 and betaglycan, can be cleaved at the plasma membrane by sheddases (from a metalloproteinase group) and this process can be triggered by PKC activation [108-110]. In order to determine whether the ACVR2 extracellular domain is shed, we designed a double-tagged receptor, with one V5 motif at the N-terminus and three consecutives HA epitopes at the C-terminus. This construct was transiently transfected in LβT2 cells, which were then treated with MG101 or the broad matrix-metalloproteinase inhibitor, GM6001, in the presence or absence of CHX. After 5 hours, the media was recovered, concentrated and immunoprecipitated with an anti-V5 antibody. As shown on Fig. 2.7C, we detected an immunoreactive ~35 kDa band in the media that could correspond to the ACVR2 glycosylated extracellular domain. The presence of GM6001 did not prevent the appearance of the band, suggesting that if shedding occurs, it is not mediated via members of the ADAM family of metalloproteases, which participates in ALK5 shedding [108]. As expected, MG101 did

82 not affect processing of this peptide (Fig. 2.7C, lanes 5 and 6); however, we detected an increase in levels of the immature receptor in the whole cell lysates (Fig. 2.7C, compare lanes 7 vs. 11). Also, we didn`t detect the presence of the 55 kDa band in the total lysates when using the V5 antibody suggesting that this fragment does not contain the N-terminal epitope. Still, this is just preliminary data that require to be further validated.

DISCUSSION

The results presented here suggest that ACVR2 is rapidly turned over in gonadotropes. We also speculate that ACVR2 might be proteolytically processed at the plasma membrane (shed); however, the exact mechanism of this cleavage remains to be elucidated.

Though we were unable to determine the half-life of endogenous ACVR2 in LβT2 cells because of a dearth of good ACVR2 antibodies, overexpressed ACVR2 decreased by 50% just 2 hours after CHX treatment. Affinity labeling/crosslinking experiments also demonstrated suppression of activin binding to LβT2 cells following 2-3 hours of CHX treatment, indicating the decrease in cell surface expression of activin type II receptors. Furthermore, knock-down experiments suggest that ACVR2 is the primary activin type II receptor in gonadotropes and that its loss cannot be compensated for ACVR2B, which is also expressed in these cells [266, 282, 299]. Consistent with this idea, genetic ablation of Acvr2 causes FSH deficiency in mice [301, 302]. Overall these data indicate that activin type II receptors, in particular ACVR2, are highly labile proteins (see also Chapters 3 and 5).

Rapid turnover of type II receptors is not unprecedented. TGFBR2 has an estimated half- life of 1 to 2.5 hours under basal conditions. In the presence of ligand (TGFβ1), its t1/2 is reduced to 1 to 1.7 hours. In contrast, the t1/2 of the type I receptor, ALK5, is considerably longer (>12h) [111-113]. Though the stability of other receptors in the family has not been formally examined, we know they are constitutively internalized even in the absence of ligand [84, 85, 87, 113] and show short residency times at the plasma membrane

83 (BMPR2 t1/2 ∼ 10 min and TGFBR2 t1/2 ∼15 min) [84, 85]. In our experiments, exogenous ACVR2 degradation was ligand-independent. Whether this reflects differential regulation of TGFBR2 and ACVR2 is not yet clear given differences in the experimental systems and cell lines used in our and previous analyses. Indeed, here we observed differential effects of CHX in different cell types. That is, CHX impaired activin A-induced SMAD2 phosphorylation in LβT2 and HaCaT, but not in HEK293 cells, suggesting that activin signaling is differentially regulated in these cell lines.

The underlying mechanisms responsible of ACVR2 fast turnover remain unresolved. Though TGFβ superfamily receptors can be degraded via either the proteasome or the lysosome [92, 98, 364, 371], none of the lysosomal inhibitors we used prevented the loss of ACVR2 in the presence or absence of CHX. Similarly, inhibition of calpains, aminopeptidases or macroautophagy failed to prevent or attenuate ACVR2 loss in CHX- treated cells, suggesting that ACVR2 may be turned over via a novel pathway.

Proteasome inhibition with MG132 partially rescued the effects of CHX on activin stimulated Fshb transcription and SMAD2 phosphorylation in LβT2 and HaCaT cells. However, in the presence of CHX, proteasome inhibitors failed to rescue the mature (or immature) receptor. Therefore, we consider that full-length ACVR2 is not degraded via the proteasome and MG132 effect on Fshb expression most likely reflects preservation of activated SMADs, which upon ligand stimulation are usually polyubiquitinated and targeted for degradation via the proteasome [92, 188, 368].

In face of the inability of proteasomal and lysosomal inhibitors to prevent ACVR2 turnover, we investigated alternative proteolytic mechanism. Different membrane receptors, including members of the TGFβ receptor family, such as ALK5, ALK3, ALK6 and BMPR2 are processed at the plasma membrane by sheddases [108-110, 372-375]. Consistent with possible ACVR2 shedding, we noted in lysates of cells treated with proteasomal inhibitors (in the absence of CHX), the appearance of a low molecular weight band (∼55 kDa), which we hypothesize is the C-terminal by product of ACVR2 proteolysis. In addition, in preliminary studies, we observed a small molecular weight

84 protein (∼35 kDa) in the cell culture media of ACVR2 expressing cells, which might correspond to the glycosylated ECD of ACVR2. In future investigations, we will determine whether ACVR2 is indeed subject to juxta-membrane and intramembrane proteolysis as well as the enzymes catalyzing these reactions.

Based on the available data, we propose that mature ACVR2 is cleaved at the plasma membrane, releasing its ECD into the media, while the intracellular fragment seems to be degraded by the proteasome, since it accumulates when this pathway is inhibited. Whether ACVR2 shedding actually occurs requires further experimentation, but if it does, most likely the cleavage is not mediated by the metalloproteinase responsible of ALK5 proteolysis.

The short half life of ACVR2 coupled with the transcriptional regulation of its gene or regulation of the shedding process itself, would allow the cell to quickly modulate its responsiveness to activins and also BMPs. In addition, ACVR2 shedding would limit activin signaling by generating a soluble decoy receptor that can both, interact with ALK4 [376] and sequester the ligand [377].

Collectively, our data indicates that the rapid turn-over of ACVR2 is responsible for the inhibin-like effects of CHX observed in gonadotropes cells. Although, we cannot rule out the possibility that the type I receptor or specific SMAD-interacting partners involved in Fshb transcription are quickly degraded as well.

85 Acknowlegments

The authors thank the indicated investigators for generously providing reagents and cell lines. This work was supported by grants from the Canadian Institutes of Health Research Grants (MOP-89991 to DJB and MOP-36379 to TEH). Carlis Rejon was supported by a scholarship from the CIHR Drug Discovery Training Program. TEH holds a Chercheur National award from the (Fonds de la Recherche en Santé du Québec (FRSQ) and DJB is a Chercheur Boursier Senior of the FRSQ. We also thank the Ligand Assay and Analysis Core at the University of Virginia Center for Research in Reproduction, which is supported by the Eunice Kennedy Shriver NICHD/NIH (SCCPIR) Grant U54-HD28934.

86 Figure legends

Figure 2.1. Prolonged exposure to cycloheximide (CHX) blocks activin A-stimulated Fshb mRNA expression in murine gonadotropes. (A) Concentration (ng/ml) of FSH and LH in spent media from pituitary primary cultures treated with different concentrations of CHX for 24 hours. Data (mean +/- SD) from 3 different experiments. Two-way ANOVA followed by Bonferroni post-hoc test. # denotes significance (p < 0.01) compared with the control condition where no CHX were added to the cells. (B) Murine primary pituitary cells (6 week-old male C57BL6/j mice) were seeded at a density of 1x105 cells/well in 96-well plates and treated with different inhibitors in 2% FBS/M199 for 24 hours. Fshb mRNA levels were determined by qRT-PCR in triplicate. Shown are the mean +/-SD of a representative experiment (n = 3). SB431542: type I receptor inhibitor; FST: follistatin-288. Statistical analysis was performed with log- transformed data. Dunnett multiple comparison test (# p < 0.01; * p < 0.05) versus the control condition. (C) Confluent LβT2 cells were incubated with CHX (5 μg/mL) for different periods of time following by a 2h treatment with activin A (25 ng/mL). RNA was isolated and Fshb mRNA levels analyzed by RT-PCR. Equal loading was confirmed by amplification of Rpl19.

Figure 2.2. CHX inhibits activin A-induced SMAD2 phosphorylation without affecting total SMAD2 protein levels in LβT2 cells. (A) LβT2 cells were serum-starved overnight and then treated with CHX (5 μg/mL) for different periods of time prior to treatment with activin A (25 ng/mL) for 1 hour. Cells were then lysed in RIPA buffer and samples analyzed by immunoblotting using the indicated antibodies. Shown is a representative blot of 4 independent experiments. (B) Quantification by densitometry (mean ± SD) of the 4 experiments performed; # p < 0.05, * p < 0.01 and $ p < 0.001.

Figure 2.3. CHX causes a time-dependent reduction in activin A binding to LβT2 cells. Confluent LβT2 cells were treated with 5 μg/mL CHX in DMEM-10% FBS. After the indicated treatment times, cells were rinsed with Krebs–Ringer-Hepes (KRH) buffer and processed as indicated in the Material and Methods section. Samples from two

87 different experiments were run on SDS-PAGE (A) while lysates obtained in other three independent experiments were quantified using a γ-counter (B). The graph shows mean (+/- SEM) of the three independent experiments with technical duplicates in each case (n=6), presented as relative specific binding (i.e. [cpm at time x/cpm in excess cold activin condition] – cpm, in excess cold activin condition). Dunnett test, * p < 0.05 compared with the time 0 condition.

Figure 2.4. ACVR2 is the primary type II receptor that mediates activin signaling in LβT2 cells. (A) LβT2 cells grown in 48-well plates were transfected with 225ng of - 1990/+1 murine Fshb-luc along with 5 nM of the indicated short interfering RNAs (siRNAs). For these experiments the Acvr2 siRNA #2 was used. Twenty-four hours after transfection, cells were treated with 25 ng/mL activin A for 24 hours. Treatments were performed in triplicate. The data represent mean +/- SEM from six independent experiments and are presented relative to the control group, in which the control siRNA was present and no ligand was added. Dunnett test * p < 0.01 (B) Fold induction (activin A versus no ligand response) for each condition. (C-D) Degradation of overexpressed ACVR2-HA is evident after 2 hours of CHX treatment. LβT2 cells transfected with 3 μg of ACVR2-HA expression vector were treated with CHX (5 μg/mL) for different period of times and cell lysates were analyzed by immunoblotting against HA or GAPDH (loading control). Representative blot of n = 3 (C) and quantification (D). Dunnett test against time t = 0h, * p < 0.01.

Figure 2.5. ACVR2 degradation is not ligand-dependent. (A-B) LβT2 cells transfected with 3 μg of ACVR2-HA or pcDNA3.1 were treated for 5 hours with CHX (5 μg/mL) in the presence or absence of activin A (25 ng/mL) and FST-288 (300 ng/mL). Cell lysates were analyzed by western blot using the indicated antibodies. Representative blot of n = 3 (A) and corresponding quantification (B). (C) LβT2 cells were transfected with 3 μg of ACVR2-HA or pcDNA3.1 and treated for 5 hours with CHX (5 μg/mL) in the presence or absence of FST-288 (300 ng/mL). Cell lysates were analyzed by western blot using the indicated antibodies. Shown is a representative blot of three independent experiments.

88 Figure 2.6. Proteasome inhibition partially reverses the effect of CHX on activin pathway. (A) LβT2 cells transfected with 3 μg of ACVR2-HA expression vector were treated with CHX for 5 hours in the presence or absence of the proteasomal inhibitor,

MG132, or the lysosomal inhibitors, leupeptin or ammonium chloride (NH4Cl). Cell lysates were analyzed by immunoblot against HA or β-actin (ACTB, as loading control). The arrow indicates the immature form of ACVR2 (B) Quantification. (C) LβT2 cells were serum-starved for 1 hour and then treated with CHX for 5 hours in the presence or absence of MG132 or leupeptin. Cells were incubated for an additional 1 hour with activin A. After treatment, cells were lysed and protein samples analyzed by immunoblotting using the indicated antibodies. (D) LβT2 cells were treated with CHX for 5 hours in the presence or absence of the proteasome inhibitor MG132. Cells were then treated for additional 2 hours with activin A (25 ng/mL). RNA was isolated, and Fshb mRNA levels were measured by qPCR. Shown are the mean +/- SD for two experiments. Dunnett test performed on log transformed data; * p <0.01.

Figure 2.7. Evaluation of possible degradation pathways for ACVR2. (A-B) Inhibition of autophagosomes, aminopeptidases or calpains has marginal effects on ACVR2 degradation. LβT2 cells transfected with ACVR2-HA construct were treated for 1 hour with 3-MA (autophagosome inhibitor) (A), MG101 (calpain inhibitor), bestatin (aminopeptidase inhibitor) or E64 (lysosomal inhibitor) (B) in serum-free media. Then CHX was added and the treatment continued for another 4 hours in the presence of the relevant inhibitors. Cell lysates were analyzed by immunoblot using α-HA or α-ACTB antibodies. Shown a representative blot of n = 3 (C) The ACVR2 extracellular domain may be shed into the media. LβT2 cells plated on 6 wells were transfected with 3 μg of V5-ACVR2-HA. The following day, cells were placed in serum-free media and treated with a broad matrix metalloprotease inhibitor (GM6001, 2.5 mM) or MG101 for 1 hour and then another 4h in the presence or absence of CHX. Conditioned media was concentrated and used for anti-V5 immunoprecipitation. The eluted product from the IPs as well as the cell lysates were analyzed by western-blot using an anti-V5 antibody. The image shown is representative of two independent experiments.

89 Supplementary Figure Legends

Supplementary Figure S2.1. CHX inhibits activin A-stimulated SMAD 2/3 phosphorylation in LβT2 cells without affecting total SMAD 2/3 levels. LβT2 cells were exposed to CHX (5 μg/mL) for different times and then treated for 1 hour with activin A (25 ng/mL). After treatment, cells were lysed in RIPA buffer and samples analyzed by immunoblotting using the indicated antibodies. The arrow indicates the band corresponding to phospho-SMAD3. The higher molecular band detected in this blot likely corresponds to phospho-SMAD1 and/or phospho-SMAD5 [305]. In the α- SMAD2/3 blot, * indicates the position of SMAD2 and ** SMAD3, which is a significantly fainter as observed previously on LβT2 cell lysates [378].

Supplementary Figure S2.2. CHX and MG132 effects on pSMAD2 levels in different cell lines. (A) MG132 partially rescues activin induced pSMAD2 levels in HaCaT and LβT2 cells. HaCaT and LβT2 cells growing on 6-well plates were treated with CHX in the presence or absence of MG132 for 5 hours, followed by 1 hour treatment with activin. Cells were lysed and whole cell lysates were analyzed by immunoblot with the indicated antibodies. Data shown are representative of two independent experiments. (B) CHX reduces basal but not activin-induced pSMAD2 levels in HEK 293 cells. Confluent HEK 293 cells were serum starved for 1 hour and then treated with CHX for 5 hours in the presence or absence of MG132. Cells were then treated for 1 hour with activin A (12.5 ng/mL), lysed and whole cell lysates were analyzed by immunoblot with the indicated antibodies. Note that two different exposure times are shown for α-SMAD2 in order to illustrate the effects of CHX on basal (not ligand present) and activin-induced levels. Data shown is from a single experiment.

Supplementary Figure S2.3. Validation of activin receptor type II siRNAs. (A) HEK- 293 cells were transfected with 1 μg of ACVR2-HA or 200 ng of ACVR2B-HA expression vectors along with 5 nM of the indicated short interfering RNAs (siRNAs). Twenty-four hours after transfection, whole cell lysates were examined by western-blot using anti-HA or anti-β-actin (ACTB, as loading control) antibodies. Representative blot

90 of two independent experiments. (B-C) LβT2 cells growing in 48-well plates were transfected with 225ng of -1990/+1 murine Fshb-luc (B) or -3740/+24 mId3-luc (C), along with 5 nM of the indicated short interfering RNAs (siRNAs). Twenty-four hours after transfection, cells were treated with 25 ng/mL activin A (B) or BMP2 (C) for 24 hours. Treatments were performed in triplicate. The data represent mean +/- SEM from six independent experiments and are presented relative to the control group, in which the control siRNA was present and no ligand was added. Dunnett test, $ represents p < 0.001.

Supplementary Figure S2.4. Effect of proteasomal inhibition on ACVR2-HA status and pSMAD2 levels in LβT2 cells. (A) Growth conditions does not significantly affect the levels of ACVR2-HA degradation. LβT2 cell were transfected with 3 μg of ACVR- HA. Twenty four hours post-transfection, cells were placed in serum-free or complete media and treated for 5 hours with CHX, in the presence of absence of proteosomal inhibitors (MG132 or epoxomycin) and subsequently lysed. Whole cell lysates were analyzed by immunoblot using the indicated antibodies. Shown is a representative blot of three independent experiments and the corresponding quantification. Bonferroni multicomparison test, * p < 0.01, $ p < 0.001. (B) ACVR2-myc is ubiquitinated. LβT2 cells were transfected with 3 μg of ACVR2-myc tagged and 0.5 μg of HA-ubiquitin. Twenty-four hours after transfection, cells were treated for 5 hours with MG132 and lysed. Fractions of whole cell lysates were used to perform anti-myc immunoprecipitation under denaturing conditions. The eluates of the IPs as well as a sample of the whole lysates were analyzed by western blot using the indicated antibodies. (C) Proteasome inhibitors attenuate the decrease in activin-induced phospho-SMAD2 levels after CHX treatment. LβT2 cells were serum starved for 1 hour and then treated for 5 hours with CHX in the presence or absence of proteasome inhibitors (10 μM MG132, 50 μM Proteasome inhibitor II (PI-II), 50 μM Proteasome inhibitor IV (PI-IV). Finally, cells were incubated with activin A or serum-free media for 1 hour before being harvested with RIPA buffer. Whole cell lysates were immunoblotted for pSMAD2. Data shown are representative of two independent experiments.

91 Supplementary figure S2.5. Lysosome inhibition does not prevent ACVR2 degradation. LβT2 cells transfected with 3 μg of ACVR2-HA or empty vector were treated for 5 hours with CHX in the presence or absence of the lysosomal inhibitors chloroquine, NH4Cl and leupeptin either alone (A) or in combination (B). Cells were harvested with RIPA buffer and cell lysates analyzed by immunoblot with anti-HA, anti- GAPDH or anti-β-actin. Data shown are representative blots of three independent experiments. (C) Testing leupeptin and NH4Cl. HEK-293 cells stable expressing a HA- tagged version of the prostaglandin F2α receptor (HEK-FP) were treated with leupeptin or NH4Cl for 5 hours in the presence or absence of CHX. Whole cell lysates were analyzed by immunoblot using the indicated antibodies. Shown here is a representative blot of two independent experiments.

Supplementary Figure S2.6. Effect of Brefeldin A on ACVR2-HA degradation in LβT2 cells. Transiently transfected LβT2 cells were incubated with the inhibitors (MG101 or CHX) 5h at 37°C in the presence or absence of brefeldin A. After treatment, cells were lysed and the protein content analyzed by immunoblot using anti-HA or anti-β- actin antibodies. The blot is representative image of three independent experiments.

92 Figure 2.1

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98 Figure 2.7

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105 Chapter 3 Activin A binds and signals via bone morphogenetic protein receptor type II (BMPR2) in immortalized gonadotrope cells

In Chapter 2, I described the rapid turnover of activin receptor type II (ACVR2) in LβT2 cells and how this might provide a mechanism for dynamic regulation of activin responses. Also, in Chapter 1, I talked about the promiscuous binding of TGFβ superfamily ligands to a comparatively finite number of receptors and how this flexibility in ligand-receptor combinations allow a single ligand to generate different responses. Here, we investigated whether activin A binds and signals via the type II receptor, BMPR2, in gonadotrope cells.

106 Activin A binds and signals via bone morphogenetic protein receptor type II (BMPR2) in immortalized gonadotrope cells

Carlis Rejón1, Mark A. Hancock2, Ying Wang1, Thomas B. Thompson3, Terence E. Hébert1 and Daniel J. Bernard1

1Department of Pharmacology and Therapeutics, and 2Sheldon Biotechnology Centre, Faculty of Medicine, McGill University, Montreal, Québec, Canada; 3Department of Molecular Genetics, Biochemistry, and Microbiology, University of Cincinnati, Cincinnati, OH, USA

Address correspondence to: Carlis Rejon, Daniel J. Bernard or Terence E. Hébert, email: [email protected], telephone: 514-398-8803, fax: 514-398-6690 email: [email protected], telephone: 514-398-2525, fax: 514-398-6690 email: [email protected], telephone: 514-398-1398, fax: 514-398-6690

107 Abstract

Regulating diverse biological processes in health and disease, TGFβ superfamily ligands greatly outnumber their receptors; therefore, receptors are shared between ligands. For example, activins and bone morphogenetic proteins (BMPs) bind and signal via the activin type II receptors ACVR2 and ACVR2. BMPs can also signal thorugh the bone morphogenetic protein type II receptor (BMPR2). We hypothesized that, in addition to its canonical receptor ACVR2, activin A can similarly bind and signal via BMPR2. First, we examined activin binding to type II receptors in heterologous cells. Radiolabeled activin A bound to ACVR2, ACVR2B and BMPR2 but not the other type II receptors (AMHR2 or TGFBR2). Using homology modeling and site-directed mutagenesis, we identified key residues in BMPR2 that mediate its interaction with activin A. In surface plasmon resonance (SPR) assays activin A bound to BMPR2 extracellular domain (ECD) with significantly lower affinity compared to ACVR2-ECD. Similarly, the ECDs of ACVR2 and BMPR2 dose-dependently inhibited activin A, but not TGFβ-induced transcriptional activity. To assess the functional significance of activin A binding to BMPR2, we altered the levels of BMPR2 expression in immortalized murine gonadotrope cells (LβT2) and evaluated the effects on activin signaling. BMPR2 over-expression potentiated activin A responses. Conversely, depletion of ACVR2 or BMPR2 with short interfering RNAs attenuated activin A-stimulated follicle-stimulating hormone β (Fshb) subunit transcription. Taken together, the data show that BMPR2 functions as a bona fide activin A type II receptor in gonadotrope cells.

108 INTRODUCTION

Transforming growth factor β (TGFβ) superfamily ligands regulate diverse biological processes, including cell proliferation and differentiation, apoptosis, organ development, wound healing, and reproduction. In general, these ligands produce their effects by binding to heterotetrameric complexes of two type I and two type II serine-threonine kinase receptors. Interestingly, the number of ligands (> 40) in the family greatly exceeds the number of available type I (7 in mammals) and type II receptors (5). Therefore, ligands bind receptors promiscuously. That is, individual receptor subtypes interact with multiple ligands and a ligand recruits different receptor combinations depending on the cellular context [41, 149, 152]. As a result, both the expression and relative affinities of different receptors determine cellular responses to the available ligands.

In pituitary gonadotrope cells, activins induce follicle-stimulating hormone (FSH) synthesis by binding to the activin type II receptors AVCR2 [301] (see also Chapter 2). The activin:ACVR2 complex then recruits a type I receptor, most often ACVR1B (or activin receptor-like kinase 4, ALK4), which in turn propagates intracellular signaling via SMAD proteins 2 and 3. SMAD2/3 partner with SMAD4 and the forkhead FOXL2 to stimulate FSH α subunit (Fshb) transcription, the rate-limiting step in FSH production [261, 300, 305, 313, 323].

Other members of the TGFβ family, specifically the bone morphogenetic proteins or BMPs (BMP2, BMP4, BMP6 and BMP7) also modulate Fshb expression [282, 325-327]. BMP2, for example, signals via BMPR2 and ALK3 in gonadotrope-like cells, to potentiate activin effects on Fshb transcription [325]. As its name suggests, BMPR2 was initially considered a BMP receptor; however, growth differentiation factors and inhibins bind BMPR2 as well [353, 379].

More recent data support the idea that activins can also interact with the BMPR2 extracellular domain (ECD) [380, 381]. Further, activin is a flexible molecule, as evidenced by the different conformations it can adopt in complex with ACVR2B [40,

109 148]. This flexibility might allow binding to other type II receptors, such as BMPR2. Notably, BMPR2 shows also conformational flexibility in its ectodomain, which could facilitate binding by different ligands [63]. Finally, both BMPs and activins, in addition to sharing type II receptors (ACVR2 and ACVR2B), bind follistatin (FST), a soluble binding protein [49, 382]. These data suggest that activins and BMPs share structural characteristics that enable their binding to common receptors.

We hypothesized that activin A binds and signals via BMPR2. As activins classically regulate FSH synthesis, we used LβT2 cells, an immortalized murine gonadotrope-like cell line, as our model system. These cells express activin receptors and BMPR2 endogenously [282], and are the standard cell line for analyses Fshb transcriptional regulation [261, 266, 282, 305, 313, 315, 323, 345, 383]. Our results show that activin A binds BMPR2 with low nanomolar affinity and signals via this receptor to regulate Fshb transcription.

MATERIALS AND METHODS

Reagents

Recombinant human ACVR2 (340-R2/CF), BMPR2 (811-BR) and TGFBR2 (341-BR) extracellular domain (ECD)-Fc chimeras, as well as human recombinant activin A, BMP2 and TGFβ1 were purchased from R&D Systems (Minneapolis, MN, USA). Enhanced Chemiluminiscence (ECL) Plus reagent and Na125I were purchased from Perkin Elmer (Waltham, MA, USA). Eagle’s minimum essential media (EMEM), 100X MEM-non- essential amino acids, 1X Dulbecco’s modified phosphate buffered-saline (D-PBS) and Dulbecco’s Modified Eagle medium (DMEM) supplemented with 4.5g/l glucose, L- glutamine (with or without sodium pyruvate) were from Wisent (St. Bruno, Quebec, Canada). DMEM/F12 with 2.5 mM L-glutamine, 15 mM HEPES buffer and 1.2 g/L sodium bicarbonate was purchased from Hyclone Laboratories (South Logan, UT, USA). Fetal bovine serum, Taq Platinum polymerase, Lipofectamine/Plus and Lipofectamine 2000 were purchased from Invitrogen Canada (Burlington, Ontario, Canada). N-

110 acetyltyrosine, urea, leupeptin, bovine serum albumin (BSA), chloramine T, potassium iodide, trichloroacetic acid (TCA), phenylmethylsulfonylfluoride (PMSF), polyethylenimine (PEI), EZview red anti-c-myc affinity gel (#E6654), EZview red anti-c- HA affinity gel (#E6779), c-myc peptide (#M2435), HA peptide (#I2149), mouse monoclonal anti-HA (#H9658), mouse monoclonal anti-myc (#M5546) antibodies were from Sigma-Aldrich Corp. (St. Louis, MO, USA). The anti-GAPDH (AM4300) antibody was purchased from Ambion (Austin, TX, USA) and horseradish peroxidase (HRP)- conjugated secondary antibodies from Bio-Rad (Hercules, CA, USA). The Nucleospin kit was from Macherey-Nagel (Düren, Germany). MMLV-RT, random hexamer primers, dNTPs and pGEM-T Easy Vector System (Cat. #A3610) were from Promega (Madison, WI, USA). Bis-sulfosuccinimidyl-suberate (BS3) was from Pierce-Thermo Scientific (Rockford, IL, USA), while G-25 sephadex PD-10 columns and the materials used for SPR experiments (CM5 sensor chip, 0.4M 1-ethyl-3-(3-dimethylpropyl)-carboiimide - EDC- in water, 0.1M N-hydroxysuccinimide -NHS- in water, 1 M ethanolamine-HCl pH 8.5 and HBS-EP buffer) were purchased from GE Healthcare (Piscataway, NJ, USA). Protein-grade detergents (Triton X-100, Empigen) were from Anatrace (Maumee, OH, USA) and Pierce Gentle Elution buffer was from Thermo Scientific (Ottawa. ON. Canada). Protease inhibitor tablets (Complete Mini) were purchased from Roche Applied Science (Laval, QC, Canada). D-luciferin potassium salt and anti-BMPR2 antibody (#612292) were obtained from BD Biosciences (Mississauga, Ontario, Canada). Oligonucleotides were synthesized by IDT (Coralville, IA, USA), which also provided the Transductin reagent. Short-interfering (si) RNAs were acquired from Dharmacon (Lafayette, CO, USA): Control (cat. # D-001210-05), Acvr2 (cat. # D-040676-01), Acvr2b (cat. # D-040629-02) and Bmpr2 (cat. # D-040599-01, 02 and 04).

Constructs

To clone the murine BMPR2 long isoform (BMPR2-L), 1 μg of LβT2 cell total RNA was reverse transcribed with MMLV-RT, random hexamer primers, and dNTPs as described previously [305]. The cDNA was subject to PCR using Taq Platinum polymerase and the following primer set: forward 5’- AGGGATGACTTCCTCGCTGC and reverse 5’-

111 CATCTCACAGACAATTCATTCCT. The PCR reaction conditions were: 94°C for 2 min, followed by 35 cycles of 94°C for 15 s, 48°C for 30 s, 68°C for 3 min and a final extension step at 68°C for 10 min. The resulting amplicon was gel-purified (Nucleospin) and TA cloned (pGEM-T Easy Vector System). Recombinant clones were identified by blue–white screening and plasmids purified (Qiaprep, Qiagen). The presence of the insert was verified by restriction digests and bidirectional DNA sequencing (Genome Québec). BMPR2-L was then subcloned into pcDNA3.0 with an in-frame C-terminal HA tag. Finally, we used QuikChange mutagenesis (Stratagene) to correct errors introduced during the initial PCR amplication (reference sequence: accession number NM007561.3). Human BMPR2 short isoform (h-BMPR2-S) and human TGFBR2 (generous gifts from Dr. J. Massague, Memorial Sloan Kettering Cancer Center, NY, USA), as well as human AMHR2-myc (provided by Dr. J. Teixeira, Massachusetts General Hospital, MA, USA), rat ACVR2-Flag and rat ACVR2B-Flag (generous gifts from Dr. T. Woodruff, Northwestern University, Chicago, IL, USA), were subcloned into pcDNA3.0 to introduce the C-terminal HA tag. Human BMPR2-ΔC-tail and human BMPR2-K230R were generated by site-directed mutagenesis of the untagged human BMPR2-S construct, by changing residue 171 from glycine to a stop codon or converting the lysine in position 230 into arginine, respectively. Human BMPR2-S-HA mutants were generated by QuikChange mutagenesis with primers indicated in Supplemental Table S3.1. Sequences of all constructs were verified by bidirectional sequencing. 3TP-luc reporter was also a gift of Dr. Massague. The ALK3-myc [325] expression vector as well as -1990/+1 murine Fshb-luc [305], -846/+1 murine Fshb-luc [311], -3740/+24 murine Id3-luc, [356] and CAGA12-luc [25] promoter-reporters were described previously.

Cell lines

Immortalized murine gonadotrope-like LβT2 cells (generously donated by Dr. P. Mellon, University of California, San Diego, CA, USA) were maintained in 10% FBS/DMEM- high glucose with sodium pyruvate. Human embryonic kidney (HEK293) cells, acquired from Invitrogen (Carlsbad, CA, USA), were grown in 10% FBS/DMEM-high glucose without sodium pyruvate. Chinese hamster (CHO) cells (a generous gift from Dr.

112 P. Morris, The Population Council, New York, NY, USA) were cultured in 10%- DMEM/F12 media. L17 cells (from Dr. J. Massague) were grown in EMEM supplemented with 10% FBS and 1X non-essential aminoacids.

Activin labelling with I125

Activin A was produced and purified as previously described [40]. This ligand was iodinated with Na125I (Perkin-Elmer, ∼17 Ci/mg) using the chloramine T method adapted from [358, 359]. Briefly, 2 μg of of activin A were diluted in 10 μl of reaction buffer (0.05M Tris, 0.15M NaCl, pH 7.4) and mixed with 1 mCi of Na125I in the presence of chloramine T solution (20 μg/ml). After 4.5 min, the reaction was stopped by addition of 20 μl 50 mM N-acetyl tyrosine, 200 μl 60mM potassium iodide and 200 μl 0.6 g/ml urea. Labeled ligand fraction was separated from unincorporated isotope using PD-10 columns equilibrated with 4 mM HCl, 75 mM NaCl, 0.1% BSA. Percent incorporation and specific activities of the proteins were determined by trichloroacetic acid precipitation. The labelled activin A had an estimated specific activity of 25-40 μCi/μg.

Binding assays and affinity labelling

CHO cells in 6-well plates were transfected when 70-80% confluent using Lipofectamine/Plus reagent and 500 ng of the indicated type II expression vectors (except for BMPR2-L, where we used 1 μg). The next day, cells were incubated 30 min at 37°C with Krebs-Ringers-Hepes (KRH) buffer pH 7.4 (128 mM NaCl, 5mM KCl, 5M MgSO4,

1.3 mM CaCl2-2H2O, 50 mM HEPES) supplemented with 0.5% BSA. Cells were then incubated for 3.5 hours with 50 pM 125I-activin A in KRH-0.5% BSA at 4°C, followed by three washes with KRH. The bound ligand was crosslinked by treatment of cells with 0.25 mM BS3 in KRH buffer (30 min incubation at 4°C). The crosslinking reaction was stopped by rinsing cells twice with 85 mM Tris pH 8; 30 mM NaCl. Cells were then harvested in 300 μl of RIPA buffer (1% NP-40, 1% deoxycholate, 0.1% SDS, 0.15M NaCl, 0.01 M sodium phosphate pH 6.8, 2 mM EDTA, 50 mM sodium fluoride and Complete Mini Protease Inhibitor Cocktail tablets) and whole cell lysates clarified by

113 centrifugation (13,000 x g, 15 min at 4°C). One hundred fifty μl of lysate were quantified with a γ counter (Packard Cobra II Auto Gamma, Perkin Elmer) or 80 μl were resolved on 8.5% acrylamide gels (running buffer: 24.8 mM Tris-HCl, 192 mM glycine, 0.1% SDS, pH 8.3) at 200 V for 4 hours at 4°C. Gels were dried for 1-2 hours at 80°C under vacuum and exposed to X-ray film.

Surface plasmon resonance (SPR)

Interactions between soluble extracellular domain receptor-Fc chimeras (85 kDa TGFBR2, 70 kDa BMPR2, 50 kDa ACVR2) and ligands (activin A; TGFβ1) or negative controls (fatty acid-free bovine serum albumin, BSA; maltose-binding protein, MBP) were examined using label-free, real-time BIACORE 3000 instrumentation (GE Healthcare Bio-Sciences AB, Uppsala, Sweden). Experiments were performed on research-grade CM5 sensor chips at 25°C using filtered (0.2 μm) and degassed HBS-EP running buffer (10 mM HEPES pH 7.4, 150 mM NaCl, 3.4 mM EDTA, 0.005% (v/v) Tween-20). Receptor extracellular domains (20 μg/ml in 10 mM sodium acetate pH 4.5) were immobilized using the Biacore Amine Coupling Kit (~900 RU final); corresponding reference surfaces were prepared in the absence of any receptor. Purified activin A (25 kDa) was injected (0 – 50 nM); over reference and receptor-immobilized surfaces using variable flow rates (25 – 50 μl/min) and contact times (5 – 10 min association/dissociation). TGFβ1 served in some experiments as positive control, and BSA or MBP were negative controls. Between sample injections, sensor chip surfaces were regenerated at 50 μl/min using two 30-second pulses of Pierce Gentle Elution buffer containing 0.1% (v/v) Triton X-100 or Empigen. Mass transport-independent data were doubled-referenced [384] and were representative of duplicate injections acquired from two independent trials. For each replicate series, a buffer blank was injected first, the highest titrant concentration second, and serial dilutions followed (from the lowest to the highest concentration repeated); comparing responses between the two highest titrant injections verified consistent immobilized surface activity throughout each assay. For

BMPR2 titrations, apparent equilibrium dissociation constants (KD) were determined by global fitting of data to a “steady-state affinity” model (BIAevaluation v4.1 software).

114 Alternatively, individual association (ka) and dissociation (kd) rate constants for the ACVR2 titrations were determined by global fitting of the data to a “1:1 kinetic” model; the inclusion of an additional mass transport coefficient did not alter the 1:1 kinetic 8 -1 -1 estimates (i.e., fitted kt parameter was ~10 RU M s as expected). Theoretical binding maxima were predicted using the following equation: Rmax = (MWA / MWL) (RL) (n);

‘Rmax’ is the maximal binding response (RU) at saturating titrant concentration, ‘MWA’ is the molecular weight (kDa) of the titrant injected in solution, ‘MWL’ is the molecular weight (kDa) of the biomolecule immobilized, ‘RL’ is the amount (RU) of biomolecule immobilized, ‘n’ is the predicted binding stoichiometry (e.g. 1:1).

Model of activin A- BMPR2binary complex

The structure of ACVR2B from a binary complex with activin A (1NYS) [40] was superimposed onto BMPR2 ECD (2HLR) [63]. To adjust the position of the ligand, activin A from 1NYS was aligned to BMP2 in 1REW. Structure figures were rendered using PyMOL [335].

Immunobloting

CHO cells in 6-well plates at 70-80% confluency were transfected using Lipofectamine/Plus reagent and 500 ng of the indicated type II expression vectors (except for BMPR2-L, where we used 1 μg). Twenty-four hours post transfection cells were washed once with 1X PBS and lysed with RIPA buffer. Cells were detached from plates using a cell scraper. Protein lysates were centrifuged 13,000 x g for 15 min at 4°C, subjected to SDS-PAGE and transferred to nitrocellulose filters (BioRad). Blots were probed with the indicated antibodies using standard techniques [305].

For siRNA experiments, HEK293 cells were cultured in 6-well plates (1x105 cell/well) and transfected 48 h later with HA-tagged receptors (200 ng of ACVR2B or 500 ng for ACVR2, BMPR2-S or BMPR2-L) along with siRNAs (5 nM final concentration) using Lipofectamine/Plus reagent. After 4-5 hours, the transfection solution was replaced with

115 growth medium and cells were allowed to recover for 18–24 hours before harvesting proteins with RIPA buffer. Whole cell protein lysates were processed as indicated above.

Gene reporter assays

LβT2 cells seeded in 48-well plates (1x105 cells/well) were transfected, when reaching 70-80% confluency, with 225 ng of reporter (-1990/+1 mFshb-luc or CAGA-luc) and various amounts of the BMPR2-L-HA expression vector or 50 ng of BMPR2-S constructs (wild type or mutant) using Lipofectamine 2000 (Invitrogen) according to manufacturer’s instructions. In each experiment, transfections were balanced with pcDNA3.1, such that all conditions included equivalent amounts of transfected DNA. In RNA interference experiments, siRNAs (5 nM final concentration) were co-transfected with the relevant reporter constructs. One day after transfection, cells were placed into serum-free media (SFM) and stimulated or not with 1 nM activin A for 24 hours.

HEK293 cells were seeded in 48-well plates and transfected with 100 ng/well of CAGA- luc reporter using Lipofectamine 2000. Twenty-four hours post-transfection, cells were cultured in serum-free media and treated or not with 5 ng/ml (~200 pM) of ligand (activin A or TGFβ1). Where indicated, ligands were incubated for 30 min at room temperature with the type II receptor extracellular domains (ECD) prior their addition to the cells. In order to test BMPR2-L functionality, L17 cells cultured in 48-well plates (4x104 cells/well) were transfected with 225 ng of 3TP-luc reporter along with 100 ng of ALK3- myc construct and/or 100 ng of either BMPR2-L or -S expression vector. DNA was balanced across treatments with pcDNA3.1 vector. The next day, cells were incubated in serum free medium or with 4 nM (100 ng/ml) of BMP2 for 24 hours.

At the end of the ligand treatment period, cells were rinsed once with 1X D-PBS and lysed in 50 μl of lysis buffer (25 mM Tris phosphate buffer pH 7.8, 10% glycerol, 1% Triton X-100, 1 mg/ml BSA and 2 mM EDTA). Luciferase activity was measured using an Orion II microplate luminometer (Berthold Detection Systems, Oak Ridge, TN, USA) using 20 μl of sample and 100 μl of assay buffer (15 mM potassium-phosphate, 25 mM

116 glycyl glycine, 15 mM MgSO4, 4 mM EGTA, 2 mM ATP, 1 mM DTT and 0.04 mM luciferin). Treatments were performed in triplicate and experiments repeated a minimum of two times.

Transfection of LβT2 cells with Transductin reagent

LβT2 cells were plated in 12 wells at ∼60% confluency. The following day, cells were placed in serum-free media and transfected according to manufacturer’s instructions using 100 μM siRNA and 1.5 μM Transductin (final concentrations). The cells were incubated for 2 hours with the transfection mixture (siRNA plus Transductin), before placing them again in complete media. After 36 hours, cells were treated for 6 hours with activin A (25 ng/ml), lysed with TRIzol reagent or RIPA buffer, to isolate RNA or protein, respectively.

Reverse transcription and quantitative PCR assays

Total RNA was extracted with Trizol reagent from LβT2 cells, according to manufacturer`s instructions. Reverse transcription was performed on 1-2 μg of total RNA. Quantitative PCR (qPCR) assays were conducted using SYBRgreen Supermix (Invitrogen) with a Corbett Rotorgene 6000 qPCR instrument (Corbett Life Science, Kirkland, Québec, Canada). Expression of Fshb was normalized relative to the gene for ribosomal protein L19 (Rpl19) and is presented relative to the “no ligand” control condition. Sequences of qPCR primers were as follows: Fshb forward, 5’- GTGCGGGCTACTGCTACACT; Fshb reverse, 5’-CAGGCAATCTTACGGTCT; Rpl19 forward, 5’-CGGGAATCCAAGAAGATTGA; Rpl19 reverse, 5’-TTCAGCTTGTG- GATGTGCTC.

Statistical analyses

Data presented are from representative or pooled experiments (as indicated in individual figures). When pooled experiments are shown (binding studies and reporter gene assays),

117 treatment replicates within an experiment were averaged to generate n=1 per treatment per experiment. The mean values from 3-6 independent experimental replicates were then used in statistical analyses such that n=3–6 per treatment. Data are presented as fold- change from control condition (no ligand and/or transfection with empty vector alone). Differences between means of untransformed or log-transformed data were compared using one- or two-way ANOVA followed by post-hoc pair-wise comparisons with Bonferroni or Dunnett adjustments as indicated (GraphPad Software, La Jolla, CA, USA). Significance was assessed relative to p < 0.05.

RESULTS

Activin binds BMPR2

We conducted SPR assays with the extracellular domains of ACVR2, BMPR2, or TGFBR2 coupled to a CM5 sensors, over which the indicated ligands were flowed over in solution. Initially, the binding specificity of each receptor-immobilized surface was screened at a fixed concentration (Supplementary Figure S3.1). In all cases, the receptor- ECD fusion surfaces were greater than 50% active (i.e., theoretical signal maximum at saturating activin A concentration, Rmax: TGFBR2 = 265 RU, BMPR2 = 321 RU, ACVR2 = 450 RU) and there was little or no binding response with BSA or MBP (i.e., negative controls), as anticipated. Activin A interacted significantly with ACVR2-ECD and BMPR2-ECD, though we observed distinctly different dissociation kinetics between the two (Supplementary Figure S3.1A-B). These interactions were specific as no activin A binding was detected when the TGFBR2-ECD was immobilized to the chip; whereas the amine-coupled receptor bound the positive TGFβ1 control ligand (Supplementary Figure S3.1C).

Activin A binding to the BMPR2- and ACVR2-immobilized surfaces was then characterized by injecting increasing concentrations of the ligand. Over equally matched surface densities (i.e. ~900 pg/mm2 of each receptor), dose-dependent and saturable binding responses were observed, but the individual kinetics and overall affinities were

118 different in each case (Table 3.1). For example, the binding of activin A to BMPR2 (Fig. 3.1A) was best characterized by a “steady-state affinity” model and the apparent equilibrium dissociation constants (KD) indicated a low nanomolar binding affinity (~14 nM). In contrast, the binding of activin A to ACVR2 (Fig. 3.1B) was best characterized by a simple “1:1 kinetic” model (i.e., the amount of complex only depended on one variable, the quantity of analyte injected) and stronger, sub-nanomolar affinity (∼0.13 nM).

Next, we performed ligand binding assays in CHO cells to confirm activin binding to BMPR2 receptor in cellulo. Two alternative spliced forms of BMPR2 have been reported, which differ by the presence or absence of a long carboxyl-terminal sequence following the intracellular protein kinase domain [9, 10]. We observed binding of 125I-activin to ACVR2 and BMPR2 (both the short –S and long isoforms –L) but not to TGFBR2 or AMHR2 (Fig. 3.1C). All five receptors were highly expressed in the cells (Supplementary Fig. S3.2). Statistical analysis verified that binding to BMPR2-S was significantly different from the control condition (vector alone). Binding to BMPR2-L was also higher than in the control condition, but did not reach significance (Fig. 3.1C, lower panel).

To gain insight in the nature of the activin A-BMPR2 interaction, we built a homology model of activin bound to BMPR2, based on the superimposition of AVCR2B-activin A (1NYS) and BMPR2-ECD (2HLR) crystal structures. The position of activin A in the complex was aligned to the structure of BMP2 (1REW) (Fig. 3.2A). From this model we predicted that three amino-acids (Y67, W85 and F115) in BMPR2 might form part of the activin A binding surface (Fig. 3.2B), while other two (H87 and Y113) would come in steric clash with activin residues (Fig. 3.2C). The accuracy of the model was evaluated by mutating those residues and performing binding assays with radiolabelled ligand.

As expected, mutations in Y67A, W85A and F115A reduced activin binding to the level seen in control cells, while the H87A slightly increased activin binding (Fig. 3.3A). Contrary to our prediction, the Y113V mutant also decreased activin binding to

119 BMPR2-S. No striking differences were detected in the total level of the mutant receptors (Fig. 3.3B); however, BMPR2-S-Y67A and BMPR2-S-F115A did not pull down efficiently in our immunoprecipitation assays, suggesting that these mutants might be conformationally distinct from the wild-type receptor. Taken together, these data indicate that activin A binds to BMPR2. Moreover, BMPR2 and ACVR2B seem to share similar activin binding surfaces.

BMPR2 extracellular domain bioneutralizes activin responses

Soluble activin type II receptors bind activins with high affinity and inhibit activin signaling by reducing ligand bioavailability in vitro [377, 385] and in vivo [386-388]. Likewise, BMPR2-ECD partially antagonized activin A-induced proliferation of granulosa cells in culture [381].

To determine whether BMPR2-ECD similarly attenuates activin signaling in other systems, we transfected HEK293 cells with the activin-TGFβ responsive reporter, CAGA-luc [25] and then treated cells with activin A or TGFβ1 in the presence or absence of increasing concentrations of ACVR2-, TGFBR2- and BMPR2-ECDs. Activin or TGFβ1 alone potently stimulated reporter activity. Pre-incubation of the ligand with either ACVR2- or BMPR2 ECDs inhibited the activin response dose-dependently (Fig. 3.4A). The ACVR2-ECD was more potent than the BMPR2-ECD, consistent with its higher binding affinity. In contrast, TGFBR2-ECD, but not BMPR2-ECD (even at the highest concentration), decreased TGFβ1-induced gene reporter activity (Fig. 3.4B). These data indicate that BMPR2 binds activin A, but not TGFβ1.

Over-expression of BMPR2-S but not BMPR2-L enhances activin A signaling in LβT2 cells

To determine the functional relevance of the activin-BMPR2 interaction, we transfected LβT2 cells with expression vectors encoding wild type (WT), ECD mutants (W85A, H87A, Y113A, H87A/Y113V) or kinase-deficient forms (K230R, ΔC-tail) of BMPR2-S

120 along with a 2 kb Fshb promoter-reporter. BMPR2-S-WT significantly increased activin A and activin B-induced Fshb-luc reporter activity (Fig. 3.5A and Supplementary Fig. S3.3). Consistent with the binding data, W85A, Y113A and H87A/Y113V mutants did not potentiate activin A signaling, whereas H87A elicited a response comparable to that of the wild-type receptor (Fig. 3.5A). Likewise, the kinase-defective (K230R) and the C- tail truncated (ΔC-tail) BMPR2-S mutants failed to increase activin-A stimulated Fshb transcription (Fig. 3.5B). In the presence of activin A, BMPR2-S also enhanced the activity of the SMAD3/4 dependent reporter, CAGA-luc, although the effect was not statistically significant (Fig. 3.5C). Collectively, the results suggest that BMPR2 can transduce activin signals in gonadotrope-like cells.

In contrast to BMPR2-S, BMPR2-L failed to potentiate activin A or activin B-stimulated Fshb-luc reporter activity (Supplementary Figs. S3.3B and S3.4), despite robust expression of the receptor in LβT2 cells (Supplementary Fig. S3.5A). BMPR2-L functionality was evaluated using two established BMPR2-dependent reporter assays. We previously showed that BMPR2-S stimulates Fshb promoter activity when over- expressed with a constitutively active form of the BMP type IA receptor (ALK3QD) in LβT2 cells [325]. Unexpectedly, here, BMPR2-L failed to simulate reporter activity under the same conditions (Fig. S3.5B). Likewise, BMP2 induces 3TP-luc reporter activity in mink epithelial cells, L17, co-transfected with BMPR2-S and BMP type I receptors [10]. Here, we similarly observed BMP2 induction of 3TP-luc in L17 cells co- transfected with either BMPR2-S or BMPR2-L (Supplementary Fig.S3.5C). These data suggest that the BMPR2-L construct (and resulting protein) is functional, but that BMPR2-S might be more biologically relevant than BMPR2-L with respect to Fshb regulation.

BMPR2 knock down inhibits activin A mediated signaling in LβT2 cells

To assess the role of endogenous BMPR2 in activin-regulated Fshb expression, we transfected LβT2 cells with siRNAs directed against Acvr2, Acvr2b or Bmpr2 followed by treatment with 1 nM activin A or activin B. In agreement with our previous results,

121 Acvr2 but not Acvr2b knockdown inhibited activin A-induced Fshb reporter promoter (Fig. 3.6A, see Chapter 2). We observed a similar pattern of results with activin B (Supplementary Fig. S3.3C). Importantly, Bmpr2 knockdown also significantly impaired activin A or B-induced Fshb-luc reporter activity (Fig. 3.6A and Supplementary Fig. S3.3C). Moreover, the activin A response was completely abrogated when cells were co- transfected simultaneously with Acvr2 and Bmpr2 siRNAs (Fig.3.6B).

Next, we examined whether activin A signals through BMPR2 to regulate endogenous Fshb transcription. In our experience, LβT2 cells transfect poorly with cationic lipids (such as Lipofectamine 2000), preventing an accurate assessment of siRNA-mediated knockdown of endogenous proteins/mRNAs [305]. Therefore, we used a newly developed peptide-based delivery system (Transductin) to introduce the siRNA into these cells. As previously reported, activin A stimulated Fshb mRNA expression [305]. This effect was significantly inhibited after transfection with Bmpr2 siRNA (Fig. 3.6C, left panel). We confirmed efficient knock-down of BMPR2 protein levels by western blot (Fig. 3.6C, right panel). These results suggest that activin A can signal via BMPR2 to induce endogenous Fshb expression in LβT2 cells.

The specificity of the Bmpr2 siRNA used in these experiments (identified as siRNA #1) was evaluated by transfecting HEK293 cells with ACVR2-HA, ACVR2B-HA or BMPR2-L-HA constructs along with control or Bmpr2 siRNA #1. Western-blot confirmed that Bmpr2 siRNA #1 reduced the expression of BMPR2-L, but not the activin type II receptors (Supplementary Fig. S3.6A). We also tested the effect of other Bmpr2 siRNAs on activin A-induced Fshb-luc reporter activity to corroborate the results obtained with Bmpr2 siRNA #1. Bmpr2 siRNA #4 diminished activin A response to a lesser extent than siRNA #1, whereas siRNA #2 did not inhibit basal or activin A- stimulated Fshb-luc reporter activity (Supplementary Fig. S3.6B). Similar results were observed with activin A stimulated CAGA-luc or BMP2-induced murine Id3-luc reporter-promoter activities (Supplementary Fig. S3.6C-D). All three siRNAs impaired expression of murine-BMPR2-L in transiently transfected HEK293 cells, but at different levels. Bmpr2 siRNAs #1 and #2 almost completely abrogated BMPR2-L expression,

122 whereas siRNA #4 reduced BMPR2-L protein levels in 75-80% and also targeted overexpressed human-BMPR2-S (Supplementary Fig. S3.6A). A fourth Bmpr2 siRNA (#3) neither knocked down BMPR2-S or BMPR-L expression nor affected the activin A response, and therefore served as an additional negative control in these experiments (Supplementary Fig. S3.6A and data not shown). These data suggest that activin A may preferentially signal via BMPR2-S compared to BMPR2-L.

Activin A regulation of Fshb via BMPR2-S seem to be ALK4-dependent

As mentioned above, BMPR2-S stimulates Fshb promoter activity when co-expressed with WT or constitutively active (QD) ALK3 [325] (Supplementary Fig. S3.5). We then examined which type I receptor complexes with BMPR2-S to mediate activin A signaling. BMPR2-S over-expression stimulated Fshb-luc reporter activity in the presence or absence of exogenous ligand (Fig. 3.7A). Treatment of cells with follistatin- 288 (FST-288), an activin bioneutralizing protein [389, 390], blocked the effect of BMPR2-S over-expression (Fig. 3.7A). As LβT2 cells synthesize activin B [282], these data suggested that endogenous activin B signaling was enhanced by BMPR2-S. FST288 also attenuated the effects of exogenous activin A in the presence or absence of over- expressed receptor (Fig. 3.7A). We observed similar results when treating cells with SB431542, a small molecule inhibitor of ALK4, 5, and 7 [204], but not BMP type I receptors (Fig. 3.7B). Given that ALK4 is the only known activin A type I receptor (i.e. ALK7 binds activin AB and B, but activin A), these data suggest that BMPR2-S’s effects on endogenous activin B and exogenous activin A are dependent on ALK4 expression.

DISCUSSION

Here, we show that activin A binds and signals via BMPR2 in LβT2 cells. Though previous studies characterized the binding properties of BMPR2 to different radiolabeled ligands in the TGFβ superfamily [9, 10, 391] and activin binding to BMPR2-ECD has been reported [380, 381], our experiments represent the first direct demonstration of an activin signaling through BMPR2.

123

A previous report indicated that BMPR2 failed to bind activin A [10]. The origin of this discrepancy is unclear, but it is probable that differences in the cell lines (COS-1 versus CHO) or transfection conditions used account for it. We tested the possibility that the cross-linkers utilized could have an impact on the detection of activin A-BMPR2 complexes, but we did not observe any difference between disuccinimidyl suberate (DSS, cell permeable) or bis-bulfosuccinimidyl-suberate (BS3, cell impermeable) in our binding assays (data not shown).

According to our SPR data, activin A binds to BMPR2 with nanomolar affinity (KD ∼14 nM). Meanwhile, fitting of the activin A-BMPR2-ECD titration series were improved using a “two-state reaction” model (BIAevaluation software, data not shown), suggesting that a conformational change might take place at the level of the receptor and/or ligand upon binding. The stronger affinity of activin A for ACVR2 (KD ∼0.13 nM) likely relates to its slower dissociation rate constant (i.e., complexes formed between ACVR2 and activin A are more stable compared to BMPR2). Physiologically, this may suggest that activin/BMPR2 interactions are more transient in nature (i.e., short-term signaling events with rapid on, rapid off kinetics) compared to activin/ACVR2 (i.e., longer-term signaling events).

The affinity of activin for BMPR2 is similar to the binding constants described for several BMPs and GDF5 [380, 392]. Moreover, since the interface that mediates BMPR2 binding to activin A seems to also be involved in BMPs and GDF5 interaction with BMPR2 (including W85 and Y113) [63, 392], our data pose the possibility that activin A and BMPs can compete for binding to BMPR2.

The functional significance of the activin-BMPR2 interaction was determined by over- expression and knockdown experiments conducted in LβT2 cells. Collectively, our data suggest a model in which activins bind to BMPR2 leading to the recruitment and phosphorylation of ALK4, which in turn induces SMAD3 activation and Fshb transcription in gonadotropes. Different observations led to this model: a) BMPR2-S

124 over-expression potentiates activin signaling, whereas kinase inactive mutants do not; b) SB451342 inhibits BMPR2-S effects on activin-induced Fshb transcription; c) over- expression of BMPR2-S also increased the activin-stimulated activity of the SMAD3/4 responsive reporter, CAGA-luc and d) transfection of Bmpr2 siRNA in LβT2 cells lead to a significant decrease in activin A-induced expression of endogenous Fshb.

Certain questions need to be addressed in order to certify the role of BMPR2 as an activin receptor. Our over-expression data suggest that activin A signals via BMPR2-S, but not the long isoform, to regulate Fshb expression in LβT2 cells. Knockdown of Bmpr2 expression with two different siRNAs (#1 and #4) decreased activin stimulated Fshb-luc and CAGA-luc reporter activities, whereas a third siRNA (#2) had no effect on activin induced responses. Given the sequences where these siRNAs anneal, we presume they target different isoforms of murine BMPR2 (noticed that we used a human-BMPR2-S expressing vector in our experiments) So, siRNA#1 and #4 presumably impair both BMPR2-S and –L forms while siRNA #2 affects only BMPR2-L expression. Therefore, future studies are needed to address whether BMPR2-S is expressed in gonadotropes and to determine the complete sequence of the murine BMPR2-S isoform (see Chapter 5).

Given that activins bind BMPR2 with weaker affinity compared to ACVR2 or ACVR2B, higher ligand concentrations and/or high expression would be expected to be important for activin signaling via BMPR2. In fact, higher expression levels of BMPR2 versus ACVR2 in LβT2 cells may be the reason why transfection of Bmpr2 siRNA reduced the Fshb-luc activity to the same extent as Acvr2 siRNA; however this hypothesis needs to be further evaluated.

In the pituitary, elevated concentrations of locally secreted activin B [287] might promote signaling through BMPR2, which is also expressed in the pituitary [282, 287, 393]. In this scenario, BMPR2 could play a role in Fshb transcriptional regulation, by transducing BMP2 [325] or activin signals. Knockdown of Bmpr2 expression in LβT2 cells actually impaired activin A stimulation of endogenous Fshb mRNA levels, although to a lesser

125 extent than the effect observed in our reporter assays. This discrepancy might be due to differences in the sensitivity of the techniques used (qPCR versus gene reporter systems).

It also remains to be established if BMPR2 can actually trans-phosphorylate ALK4 and transduce activin signals by itself or if recruitment of ACVR2 to the receptor complex is required. In preliminary analyses, we observed dimerization of BMPR2 and ACVR2 (data not shown). However, the interacting proteins lacked mature glycosylation patterns, suggesting that the interaction was occurring intracellularly, perhaps in the ER. Moreover, we have been unable to detect interaction of the receptors at the plasma membrane. Thus, BMPR2 might interact directly with ALK4 in the absence of ACVR2.

Overall, our results indicate that BMPR2 is a low affinity activin type II receptor in LβT2 cells. Therefore, expression levels of ACVR2, ACVR2B (i.e. higher affinity receptors) and BMPR2 will determine cell sensitivity to activin stimulation and possibly to the activation of distinct canonical and alternative pathways. On the other hand, activin expression might affect BMPs and GDFs association to BMPR2, because these ligands seem to share the same binding interface. Finally, the potential interaction between ALK4 and BMPR2 needs to be further characterized.

Acknowlegments

The authors thank the indicated investigators for generously providing reagents and cell lines. This work was supported by grants from the Canadian Institutes of Health Research Grants (MOP-89991 to DJB and MOP-36379 to TEH). Carlis Rejon was supported by a scholarship from the CIHR Drug Discovery Training Program. TEH holds a Chercheur National award from the (Fonds de la Recherche en Santé du Québec (FRSQ) and DJB is a Chercheur Boursier Senior of the FRSQ.

126 Figure Legends

Figure 3.1. Activin A binds to BMPR2. (A-B) Surface Plasmon resonance experiments. Label-free, real-time kinetics of activin A (0-12.5 or 50 nM; 2-fold serial dilutions) binding to amine-coupled receptor extracellular domains (~900 RU each; 50 μL/min x 10 min association + 10 min dissociation). (A) Representative BMPR2 ECD data (solid lines) were fit globally to a steady-state affinity model (inset). (B) Representative ACVR2 ECD data (solid lines) were fit globally to a 1:1 kinetic model (dashed lines). (C) Binding assays. CHO cells transfected with 0.5 μg (ACVR2, BMPR2-S, TGFBR2, AMHR2) or 1 μg (BMPR2-L) of HA-tagged receptor expression vector. One day after transfection, cells were affinity labelled by incubation with 50 pM of I125-activin A, followed by crosslinking with bis-sulfosuccinimidyl-suberate (BS3). Cell lysates were either subjected to SDS-PAGE and autoradiography (top) or quantified using a γ-counter (bottom). Data represent mean (+/-SEM) of three independent experiments and are presented relative to the cells transfected with pcDNA3.1. Dunnett comparison test against pcDNA3.1 condition, # p < 0.01; * p <0.05.

Figure 3.2. A hypothetical model of activin A bound to BMPR2. (A) The homology model was created by superimposing the structure of BMPR2 (2HLR, magenta) to the one of ACVR2B (cyan) bound to activin A (1NYS). At the same time activin A (green) was aligned to the position of BMP2 when bound to ALK3 (1REW, not shown). Highlighted in yellow are three of the residues that were evaluated in our study (see text). (B) Residues predicted to favour activin A binding to BMPR2. (C) Residues predicted to preclude activin A binding to BMPR2.

Figure 3.3. BMPR2 mutants affect activin A binding. (A) Activin A binding. CHO cells seeded in 6-well plates were transfected with 0.5 μg of wild-type or mutant HA- tagged BMPR2-S expression vectors. The next day, cells were affinity labelled as indicated in Materials and Methods. Cell lysates were quantified on a γ-counter. Data are the mean (+/-SEM) of three independent experiments and are presented relative to the cells transfected with pcDNA3.1. Dunnett comparison test against pcDNA3.1 condition,

127 # p < 0.01. (B) Expression of BMPR2 mutants. CHO cells were transfected as indicated above. Twenty-four hours after transfection cell surface proteins were biotinylated and whole cell lysates collected for HA immunoprecipitation analysis. Immunoprecipitated proteins were immunoblotted (IB) with HA antibody or using ABC Vectastain reagent to detect biotinylated receptors. GAPDH was used a loading control.

Figure 3.4. BMPR2 ECD antagonizes activin A but not TGFβ1 signaling. HEK293 cells plated in 48-well plates were transfected with CAGA-luc reporter (100 ng/well). The next day, cells were treated with 5 ng/ml activin A (A) or TGFβ1 (B) in the presence or absence of recombinant receptor extracellular domains (ECD). Ligands were pre- incubated 30 min with ECDs at room temperature, prior to addition to the cells. The “No ligand” condition refers to cells placed in serum-free media. Cells were treated for 24 hours before samples were processed. Treatments were conducted in triplicate. Data are means (+/-SD) of two independent experiments (n=2). Statistical analysis was done with log transformed data. Dunnett comparison test against “Activin A alone” or “TGFβ1 alone” conditions, # p < 0.01; * p <0.05.

Figure 3.5. Over-expression of BMPR2-S enhances activin A signaling in LβT2 cells. LβT2 cells seeded in 48 well plates were transfected with murine -1990/+1 Fshb-luc (A- B) or CAGA-luc (C) reporter (225 ng/well) along with 50 ng wild-type (WT) or mutant BMPR2-S expression vectors. One day post-transfection, cells were treated with 25 ng/ml activin A in serum-free medium for 24 hours. In (A), different ECD mutants were tested, while in (B-C) two different BMPR2-S kinase impaired mutants were used. One day post-transfection, cells were treated with 25 ng/ml activin A in serum-free medium for 24 hours. Treatments were performed in triplicate. Data represent mean (+/-SEM) of three- four independent experiments. Values were normalized relative to the control group in which cells were transfected with pcDNA3.1 and no ligand was added. Bonferroni multicomparison test, $ p < 0.001; # p < 0.01 * p < 0.05.

Figure 3.6. BMPR2 knockdown attenuates activin A-mediated signaling in LβT2 cells. LβT2 cells seeded in 48-well plates were transfected with murine -1990/+1 Fshb-

128 luc reporter (225ng/well) and 5 nM (final concentration) of control, Acvr2, Acvr2b, or Bmpr2 short interference RNA (siRNA) alone (A) or in combination (B). In the combined condition 2.5 nM of each siRNA was used. Cells were treated with 25 ng/ml activin A in serum-free medium for 24 hours. Treatments were performed in triplicate. Data represent the mean (+/-SEM) of six independent experiments. Values were normalized relative to the control group in which control siRNA was transfected and no ligand was added. Bonferroni comparison test, * p < 0.05; $ p < 0.001 compared to corresponding control condition. (C) LβT2 cells plated on 6 wells at 80% confluency were transduced with control or Bmpr2 siRNA #1 (100 μM) in the presence of Transductin (1.5 μM). Thirty-six hours post-transduction, the cells were treated for 6 hours with activin A (25 ng/ml), lysed and the RNA or proteins were extracted. In the left panel is shown the change in Fshb expression measured by qPCR. Values were normalized to the housekeeping gene, Rpl19 and are presented relative to the control condition were no ligand was added. The data correspond to the mean (+/-SEM) of three independent experiments. Bonferroni post-test, * p < 0.05. To the right is a representative blot showing the knockdown of the endogenous receptor in cells transducted with Bmpr2 siRNA. An anti-β-actin antibody was used as loading control.

Figure 3.7. Inhibitors of activin signaling counteract effects of BMPR2-S over- expression. LβT2 cells seeded in 48-well plates were transfected with 225 ng/well of murine -1990/+1 Fshb-luc along with 50 ng WT-BMPR2-S expression vector. (A) One day post-transfection, cells were treated with either 25 ng/ml activin A alone, activin A preincubated for 30 minutes with 300 ng/ml FST-288 or FST-288 alone for 24 hours. In (B) cells were exposed to 10 μM SB431542 in serum free medium for 30 minutes and then treated (were indicated) with 25 ng/ml activin A in the continued presence of the inhibitors for 24 hours. Treatments were performed in triplicate. Data represent the mean (+/-SEM) of three independent experiments. Values were normalized relative to the control group in which cells were transfected with pcDNA3.1 and no ligand was added. Bonferroni comparison test, $ p < 0.001.

129 Supplementary Figure legends

Supplementary Figure S3.1. Activin A binds to ACVR2 and BMPR2 but not TGFBR2 extracellular domains. Before performing the real-time kinetic experiments, the interaction of positive and negative controls to the amine coupled receptor extracellular domains was assessed by SPR (~900 RU each; 25 μL/min x 5 min association + 5 min dissociation). (A) ACVR2-ECD; (B) BMPR2-ECD; (C) TGFBR2- ECD. Activin A (50 nM) was used as a positive control for ACVR2-ECD surface, and TGFβ1 (50 nM) was the control for TGFBR2. BSA and MBP (50 nM) were used as negative controls (data not shown).

Supplementary Figure S3.2. Expression levels of HA-tagged receptors. CHO cells seeded in 6-well plates were transfected with 0.5 μg (ACVR2, BMPR2-S, TGFBR2, AMHR2) or 1 μg (BMPR2-L) of HA tagged receptor or pcDNA3.1 and harvested 24h post-transfection. Lysates were blotted with HA antibody. The cells were transfected in parallel with those used in binding assays (Figure 3.1C). L, long isoform; S, short isoform.

Supplementary Figure S3.3 Activin B signaling is also affected by BMPR2-S expression in LβT2 cells. (A-B) Activin B signals though BMPR2-S receptor. LβT2 cells seeded in 48-well plates were transfected with the murine -1990/+1 Fshb-luc reporter (225 ng/well) along with BMPR2-S, ACVR2, ACVR2B (50 ng/well) or BMPR2-L (100 ng/well) expression vectors. One day post-transfection, cells were treated with 25 ng/ml activin B in serum-free medium for 24 hours. Treatments were performed in triplicate. Data represent mean (+/-SEM) of three independent experiments. Values were normalized relative to the control group in which cells were transfected with pcDNA3.1 and no ligand was added. Bonferroni comparison test, # p < 0.01; $ p < 0.001. (C) LβT2 cells seeded in 48-well plates were transfected with the murine -1990/+1 Fshb-luc reporter (225 ng/well) and the indicated siRNAs (5 nM final concentration). Cells were treated with 25 ng/ml of activin B in serum-free medium for 24 hours. Treatments were performed in triplicate. Data represent mean (+/-SEM) of 3-4 independent experiments.

130 Values were normalized relative to the control group in which control siRNA was transfected and no ligand was added. Bonferroni multicomparison test, * p < 0.05; # p < 0.01; $ p < 0.001.

Supplementary Figure S3.4. Over-expression of BMPR2 long isoform does not enhance activin A signaling. LβT2 cells seeded in 48-well plates were transfected with murine -1990/+1 Fshb-luc reporter (225 ng/well) and increasing amounts of BMPR2-L expression vector. One day post-transfection, cells were treated with 25 ng/ml activin A in serum-free medium for 24 hours. Treatments were performed in triplicate. Data represent mean (+/-SEM) of three independent experiments. Values were normalized relative to the control group in which cells were transfected with pcDNA3.1 and no ligand was added

Supplementary Figure S3.5. The BMPR2-L construct is expressed and functional. (A) LβT2 cells seeded in 6-well plates were transfected with 3 μg of BMPR2-L (untagged, used as negative control or HA-tagged) expression vectors and lysed 24 h later. Whole protein cells lysates were collected and subjected to immunoblot analysis with HA antibody. The arrow indicates the bands corresponding to the HA tagged BMPR2-L. (B) LβT2 cells seeded in 48-well plates were transfected with murine -846/+1 Fshb-luc (250 ng/well) along with 50 ng/well pcDNA3.1 or a constitutively active ALK3 receptor expression vector (ALK3QD). Cells were co-transfected with 50 ng BMPR2-S or BMPR2-L HA-tagged receptors. One day after transfection, cells were placed in serum-free media and incubated for 24 hours. Treatments were performed in triplicate. The figure shown is from a representative experiment of three independent assays (C) L17 cells seeded in 48-well plates were transfected with 225 ng/well of 3TP-luc reporter along with 50 ng/well of the indicated receptor expression vector. The conditions were balanced for total DNA with pcDNA3.1. The next day, cells were treated with BMP2 (25 ng/ml) in serum-free media for 24 hours. Treatments were performed in triplicate. The figure shown is from a representative experiment of two independent assays.

131 Supplementary Figure S3.6. Evaluation of Bmpr2 siRNAs. (A) Knock-down of overexpressed receptors in HEK293 cells. HEK293 cells seeded in 6-well plates were transfected with 200 ng (rat ACVR2B) or 500 ng (rat ACVR2, human BMPR2-S, murine BMPR2-L) of HA-tagged type II expression constructs along with 5 nM (final concentration) of the indicated siRNAs. The next day, cells were lysed and samples used for immunoblot analysis with HA antibody. GAPDH was used as a loading control. (B- D) Gene reporter assays. LβT2 cells seeded in 48-well plates were transfected with murine -1990/+1 Fshb-luc (B), CAGA-luc (C) or murine -3470/+24 Id3-luc (D) reporters (225 ng/well) and the indicated siRNAs (5 nM final concentration). Cells were treated with either 25 ng/ml of activin A or BMP2 in serum-free medium for 24 hours. Treatments were performed in triplicate. Data represent mean (+/-SEM) of three to six independent experiments. Values were normalized relative to the control group in which control siRNA was transfected and no ligand was added. Bonferroni comparison test, * p < 0.05; # p < 0.01; $ p < 0.001.

132 Figure 3.1 A C

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133 Figure 3.2

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134 Figure 3.3

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135 Figure 3.4

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136 Figure 3.5

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137 Figure 3.6

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138 Figure 3.7

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139 Supplementary Figure S3.1

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140 Supplementary Figure S3.2

141 Supplementary Figure S3.3

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142 Supplementary Figure S3.4

143 Supplementary Figure S3.5 A C

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144 Supplementary Figure S3.6

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145 Table 3.1: Apparent kinetics of activin A binding to amine-coupled receptor-Fc fusions, as assessed by surface plasmon resonance (SPR) 5 -1 -1 -4 -1 Immobilized Surface ka x 10 M s kd x 10 s KD (nM) BMPR2a 4.7 +/- 0.02 85 +/- 0.5 14 +/- 1 ACVR2b 44 +/- 0.01 5.6 +/- 0.03 0.13 +/- 0.02 a Equilibrium dissociation constants (KD) represent global analysis of titration series using a “steady-state affinity” model (+/-SEM; n=2); individual association (ka) and dissociation (kd) rate constants were determined using the “Fit Separate ka / kd” tool. b Kinetic estimates represent global analysis of titration series to a “1:1 kinetic” model 8 -1 -1 (+/- SEM; n=2) and were independent of mass transport limitations (kt ~10 RU M s ).

146 Supplementary Table S3.1: Primers used for BMPR2-S mutagenesis† hBMPR2-Y67A-FWD TCGAAAGGTAGCACCTGCgcTGGCCTTTGGGAGAAATC hBMPR2-W85A-FWD ATCTTGTAAAACAAGGATGTgcGTCTCACATTGGAGATCCCC hBMPR2-H87A-FWD AAAACAAGGATGTTGGTCTgcCATTGGAGATCCCCAAGAG hBMPR2-Y113V-FWD TCCTCCCTCAATTCAGAATGGAACAgtCCGTTTCTGCTGT hBMPR2-F115A-FWD CAATTCAGAATGGAACATACCGTgcCTGCTGTTGTAGCACAGATTTAT

† Only the sense strand is shown. Mutations are shown in lowercase.

147 Chapter 4 Preliminary assessment of TGFβ type I receptor combinatorial interactions using Bioluminescence Resonance Energy Transfer (BRET) assays

In the previous two chapters, I showed novel examples that illustrate how signaling in the TGFβ family (activin specifically) is modulated by rapid receptor turnover (Chapter 2) or novel receptor-ligand interactions (Chapter 3). I also noted in Chapter 1 how signaling diversity can be achieved through different receptor combinations, using as an example TGFβ signaling trough ALK5 or ALK1-ALK5 heterodimers in endothelial cells. In this chapter, I conducted a preliminary screen of potential type I receptor dimer combinations, using bioluminescence resonance energy transfer (BRET), a biophysical approach previously validated in our laboratory to study G protein-coupled receptor signaling complexes.

148 Preliminary assessment of TGFβ type I receptors combinatorial interactions using Bioluminescence Resonance Energy Transfer (BRET) assays

Carlis Rejón*, Irina Glazkova, Eugénie Goupil, Daniel J. Bernard* and Terence E. Hébert*

Department of Pharmacology and Therapeutics, Faculty of Medicine, McGill University, Montreal, Québec, Canada; 3655 Promenade Sir William Osler, Montreal Quebec, H3G 1Y6 Canada

*Address correspondence to: Carlis Rejon, Daniel J. Bernard or Terence E. Hébert, email: [email protected], telephone: 514-398-8803, fax: 514-398-6690 email: [email protected], telephone: 514-398-2525, fax: 514-398-6690 email: [email protected], telephone: 514-398-1398, fax: 514-398-6690

149 Abstract

Members of the TGFβ superfamily signal via heterotetrameric complexes of two type I and two type II transmembrane serine/threonine kinase receptors. Variations in the composition of these receptor complexes contribute to the generation of signaling diversity. To date, seven type I and five type II receptors have been characterized in mammals. However, out of the 21 theoretically possible type I receptor hetero- and homo-dimer combinations only five have been confirmed to date. Given the predicted structural similarity among the type I receptors, one might envision that more heterodimers might be possible beyond those currently reported. We conducted a preliminary assessment of the combinatorial possibilities for heterodimeric type I receptor interactions using bioluminescence resonance energy transfer (BRET). We observed significant promiscuity among the type I receptors, particularly for activin receptor-like kinase 4 (ALK4). On the other hand, ALK5 seemed more likely to dimerize with itself. Unexpectedly, ALK4 and ALK5 were also capable of interacting with β2- adrenergic receptor, as determined in preliminary BRET and receptor co-precipitation experiments. Greater flexibility in terms of type I receptor heterodimerization would further contribute to signaling diversity in the TGFβ family. Moreover, potential crosstalk between ALK4 and ALK5 with β2AR may have implications in function and remodelling during the progression of heart disease.

150 INTRODUCTION

The TGFβ superfamily of ligands encompasses a large group (> 30) of structurally- related dimeric proteins [2]. These ligands mediate a variety of cellular functions, and dysregulation of their signaling is implicated in several diseases including cancer, fibrosis, skeletal malformations, infertility and heart failure [6, 7, 394]. All TGFβ family members signal via complexes of two type I and two type II transmembrane serine/threonine kinase receptors [10, 11, 13-15] and the final response elicited critically depends on the combination of receptors present in the signaling complex, likely in a cell type-specific manner.

Type I and type II receptors show considerable sequence homology, although the sequence similarity within the type I receptor sub-family is more striking [65, 66]. Both, type I (known as activin receptor-like kinases or ALKs) and type II receptors share common structural features, such as the presence of a cysteine-rich extracellular domain, a single-pass transmembrane domain, and a large intracellular Ser/Thr protein kinase domain [3, 66]. In contrast to the large number of ligands, in vertebrates there are only five type II and seven type I (ALK1-7) receptors, therefore there is significant overlap in receptor usage between ligands. However, distinct type I-type II receptor combinations allow differential ligand binding and generate signaling diversity [180].

Although type I and type II receptors were originally regarded as homo-oligomers [66, 395], a few hetero-dimeric combinations have been reported among type I receptors [121, 130, 131, 137-139, 396]. However, given the predicted structural resemblance between type I receptors, these novel combinations only constitute a minor subset of theoretically possible dimeric associations [39, 62, 64]. This might reflect limitations in the analyses conducted to date and/or biological restraints that may prevent the association of certain receptors. Moreover, at present the key determinants driving type I receptor dimerization are poorly understood. In fact, a systematic evaluation of the combinatorial possibilities among the different type I receptors has never been reported or has been interrogated

151 with a limited number of approaches. Here, we explored the potential of type I receptor heterodimerization using bioluminescence resonance energy transfer (BRET) analysis.

We discovered that there is considerable promiscuity among type I receptor interactions; ALK1 and ALK4 seem more prone to hetero-dimerization with other members of the family, whereas ALK5 was more likely to homodimerize, indicating that our approach presents certain limitations as it was unable to detect ALK5 heterodimers already reported in the literature. Surprisingly, we also found that the β2-adrenergic receptor (a G protein-coupled receptor, GPCR) interacted with ALK4 and ALK5, using BRET and co- precipitation assays.

MATERIALS AND METHODS

Reagents

Human recombinant activin A and TGFβ1 were purchased from R&D Systems (Minneapolis, MN, USA). Enhanced Chemiluminescence (ECL) Plus reagent was purchased from Perkin Elmer (Waltham, MA, USA). Fetal bovine serum (FBS), 1X Dulbecco’s modified phosphate buffer solution (D-PBS) and Dulbecco’s Modified Eagle medium (DMEM) supplemented with 4.5g/l glucose, L-Glutamine were from Wisent (St. Bruno, Quebec, Canada). EZview red anti-c-myc affinity gel (#E6654), c-myc peptide (#M2435), horseradish peroxidase (HRP)-conjugated secondary antibodies, polyethylenimine (PEI) and all the reagents used for crude membrane preparation (unless otherwise stated) were purchased from Sigma-Aldrich Corporation (St. Louis, MO, USA). The anti-GAPDH (AM4300) antibody was purchased from Ambion (Austin, TX, USA). The mouse monoclonal antibodies anti-HA (MMS-101R) and anti-myc (MMS- 150), produced by Covance, were acquired from Cederlane (Hornby, ON, Canada). Protease inhibitor tablets (Complete Mini) were purchased from Roche Applied Science (Laval, QC, Canada), puromycin from InvivoGen (San Diego, CA, USA) and streptavidin-sepharose beads from GE Healthcare (Uppsala, Sweden). Coelenterazine

152 400A was obtained from Biotium (Hayward, CA, USA). Lipofectamine 2000 was purchased from Invitrogen Canada (Burlington, Ontario, Canada).

Constructs

Human TGFBR2, human BMPR2, human ALK1-HA and 3TP-luc reporter were gifts from Dr. Joan Massague (Memorial Sloan Kettering Cancer Center, NY, USA). The rat ALK4-HA and rat ALK2-HA expression vectors were generously provided by Dr. T.

Woodruff (Northwestern University, Chicago, IL, USA). β2AR-Rluc and β2AR-GFP10 constructs were previously described [397]. Rat ALK5-FLAG in pRK5F [398] was a kind gift of Dr. Y. Zhang (National Cancer Institute, Bethesda, MD, USA.). The CD8-Rluc [399], human ALK3-myc [325] constructs as well as -1990/+1 murine Fshb-luc [305] and CAGA12-luc [25] promoter-reporters were described previously. The TAP-β2AR, which expresses a β2AR receptor fused to a TAP tag (containing streptavidin- and calmodulin-binding epitopes and hemaglutinin -HA- tag) at its N-terminus was generated in the laboratory and verified by sequencing [365]. For BRET experiments, the type I receptors ALK1-HA, ALK2-HA, ALK3-myc, ALK4-HA and ALK5-FLAG were subcloned into humanized pRluc-N and p-GFP2-N vectors (Perkin-Elmer, Montréal, QC, Canada), so that the Rluc or GFP2 tags are located at the C-terminus of the receptors. To conduct protein fragment complementation assays (PCA), we used the ALK5- and β2AR- split Venus constructs previously described [400, 401]. These vectors render a protein tagged at its C-terminus with the N-terminal portion (residues 1-158) or the C-terminus (amino acids 159-239) region of the yellow fluorescent protein variant Venus. ALK1,

ALK2, ALK4, ALK5 and β2AR, were also subcloned into pcDNA4/myc-His vector (Invitrogen) to create myc-tagged version of these receptors.

Cell lines

Human embryonic kidney (HEK293) cells acquired from Invitrogen (Carlsbad, CA, USA) were grown in 10% FBS/DMEM-high glucose without sodium pyruvate. HEK293 cells stably expressing the TAP-β2AR, were previously generated and analyzed in the

153 laboratory [365]. These cells (identified as HEK-TAP-β2AR) were grown in 10% FBS/DMEM-high glucose without sodium pyruvate and selected with 1 μg/mL puromycin. Mink lung epithelial derived cell lines, L17 and DR26 cells (from Dr. J. Massague), were grown in EMEM supplemented with 10% FBS and 1X non-essential aminoacids. Immortalized murine gonadotrope LβT2 cells (generously donated by Dr. P. Mellon, University of California, San Diego, CA, USA) were maintained in 10% FBS/DMEM-high glucose with sodium pyruvate.

Gene reporter assays

LβT2 cells seeded in 24-well plates (2x105 cells/well) were transfected 2 days later with 450 ng of -1990/+1 mFshb-luc along with 100 ng of receptor expression vectors using Lipofectamine 2000 (Invitrogen) according to manufacturer’s instructions. In each experiment, transfections were balanced with pcDNA3.1, such that all conditions included equivalent amounts of transfected DNA. In ALK1, ALK2 and ALK3 gene reporter assays, BMPR2 (100 ng/well) was co-transfected. One day after transfection, cells were placed into serum-free media (SFM). To test ALK4-myc functionality, cells were stimulated or not with 25 ng/ml activin A for 24 hours.

L17 and DR26 cells cultured in 24-well plates (1-2x105 cells/well) were transfected with 450 ng reporter (3TP-luc or CAGA-luc) along with 100 ng TGFBR2-Rluc, 100 ng ALK5-Rluc or 50 ng ALK5-myc constructs, using Lipofetamine 2000. DNA was balanced across treatments with pcDNA3.1. The next day, cells were incubated in serum free medium with or without 5 ng/ml of TGFβ1 for 24 hours.

At the end of the treatment period, cells were rinsed once with 1X D-PBS and lysed in 50 μl lysis buffer (25 mM Tris phosphate pH 7.8, 10% glycerol, 1% Triton-X100, 1 mg/ml BSA and 2 mM EDTA). Luciferase activity was measured using an Orion II microplate luminometer (Berthold Detection Systems, Oak Ridge, TN, USA) using 20 μl sample and 100 μl assay buffer (15 mM potassium-phosphate, 25 mM glycyl glycine, 15 mM

154 MgSO4, 4 mM EGTA, 2 mM ATP, 1 mM DTT and 0.04 mM luciferin). Treatments were performed in duplicate or triplicate.

Bioluminescence resonance energy transfer (BRET)

BRET assays were performed as previously described [399, 402]. Briefly, HEK293 cells were seeded into 6-well plates (1.5-2x105 cell/well) and transfected 2 days later with PEI (Sigma; 1 mg/ml stock) at 1:3 ratio with DNA. In the saturation assays, HEK293 cells were transfected with 0.2 μg (ALK5, TGFBR2, or CD8) or 0.1 μg (β2AR) of donor (Rluc-tagged) and increasing amounts (0.125, 0.25; 0.5; 1 and 2 μg) of BRET acceptor (GFP2-conjugated) partner in 1 ml serum free medium (SFM). Total DNA amount was balanced in all conditions by adding pcDNA 3.1 as required. After 4 hours incubation, 1 ml of complete media was added per well and cells were allowed to recover for 48 hours. For the homo- and hetero-dimerization screening of type I receptors, only three concentrations of acceptor-expressing (GFP2) vectors were used (0.5, 1 and 1.5 μg of DNA) together with 0.5 μg Rluc-tagged protein. In the competition assays, the HEK293 cells were transfected with 0.4 μg ALK5-Rluc, 1 μg ALK5-GFP2 along with two different concentrations (0.75, 1.5 or 3.0 μg) of different myc-tagged type I receptor

(ALK1, ALK2, ALK3, ALK4 or ALK5) or myc-β2AR-expressing vectors. Again, total amount of DNA was balanced with pcDNA3.1.

Forty-eight hours after transfection, cells were washed once with 1X PBS, harvested in 500 μl of the same buffer and allowed to settle by gravity. The cell pellet (in 90 μl of PBS) was loaded into white 96-well microplates (Optiplate, PerkinElmer Life and Analytical Sciences). BRET assays were conducted using coelenterazine 400A (10 μl) as substrate at a final concentration of 5 μM. Signals were collected on a Packard Fusion instrument (PerkinElmer Life and Analytical Sciences) using 410/80-nm (luciferase- donor) and 515/30-nm (GFP-acceptor) band pass filters. BRET ratios were calculated by dividing the amount of the light passed by the 515/30-nm filter to that passed by the 410/80-nm filter (acceptor emission/donor emission). To test the effect of the TGFβ1 ligand in ALK5 oligomerization, cells were incubated 20 min at 37C in serum free media

155 with or without TGFβ1 (5 ng/mL), before being rinsed and detached for BRET experiments.

Cell membrane preparations

One T75 flask with confluent HEK-TAP-β2AR cells was transiently transfected with 10 μg of pcDNA3.1, ALK5-myc or other indicated type I receptor expression vectors using Lipofectamine 2000, according to the manufacturer’s instructions. Forty eight hours later, total crude membrane were prepared from cells as previously described [365]. In brief, cells were rinsed once with PBS, pelleted and disrupted with 10 ml of lysis buffer (5 mM Tris-HCl pH 7.4, 2 mM EDTA, 5 μg/ml trypsin inhibitor, 10 μg/ml benzamidine, 5 μg/ml leupeptin). Lysates were homogenized with a Polytron (Ultra Turrax T18 basic, IKA) and centrifuged 1000 x g for 5 min at 4°C. The supernatant was recovered and ultracentrifuged (30,600 x g for 20 min at 4°C) to obtain crude membranes. These crude membrane preparations were resuspended overnight at 4°C on a rocking platform with 1 ml of solubilization buffer [75 mM Tris-HCl pH 8.0, 2 mM EDTA, 5 mM MgCl2, 0.5% n-dodecyl β-D-maltoside (DDM), 5 μg/ml trypsin inhibitor, 10 μg/ml benzamidine, 5 μg/ml leupeptin]. The next day, samples were centrifuged for 20 min at 40,000 x g. A fraction (25-50 μl) of the solubilised crude extract was reserved (total lysates) before proceeding with the streptavidin bead purification or immunoprecipitation (see below).

Streptavidin bead purification

Solubilized membrane preparations were incubated with 100 μl of pre-equilibrated streptavidin-Sepharose beads overnight at 4°C on a rocker. The following day, beads were washed four times with 500 μl solubilization buffer (75 mM Tris-HCl pH 8.0, 2 mM

EDTA, 5 mM MgCl2, 0.5% DDM, 5 μg/ml trypsin inhibitor, 10 μg/ml benzamidine, 5 μg/ml leupeptin). Proteins were eluted by resuspending the beads in 50 μl of 4X SDS PAGE loading buffer (250 mM Tris, pH 6.8, 8% SDS, 40% glycerol, 0.2% bromophenol blue, 5% β-mercaptoethanol) and heating them for 15 min at 65°C before analyzing the samples by SDS-PAGE and immunoblot.

156

Immunoprecipitation (IP)

Solubilized membrane preparations obtained from stable TAP-β2AR HEK293 cells transiently transfected with ALK5-myc or pcDNA3.1 were subjected to immunoprecipitation using EZ view Red anti-myc beads following manufacturer instructions. In brief, 900 μl protein lysate were incubated overnight with 50 μl equilibrated beads on a rotating platform at 4°C. After three washes with lysis buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100), the beads were incubated with 50 μl of c-myc peptide solution (30 min at 4°C) on a rotating platform to elute gel-bound proteins which were then analyzed by immunoblot.

Immunoblotting

Total protein lysates from TAP-β2AR transiently transfected cells and protein eluates obtained after streptavidin bead purification or immunoprecipitation protocols were subjected to SDS-PAGE using 10% acrylamide gels and transferred onto activated PVDF membranes (BioRad). Membranes were blocked with 5% milk dissolved in TBS-0.1% Tween solution and probed with the indicated antibodies using standard techniques [305].

Protein fragment complementation assay and confocal microscopy

HEK293 seeded in 6 well plates (1x105 cells/well) were transfected with equal amounts of ALK5 and/or β2AR split Venus constructs (as indicated) using a conventional calcium phosphate co-precipitation protocol. Forty eight hours after transfection, cells were cultured in serum free medium for 30 min. Images were collected using a Zeiss LSM-510 system (Carl Zeiss MicroImaging Inc, Thornwood, NY) using excitation at 515 nm for YFP and emission measured with the BP530-600 filter set.

157 Statistical Analysis

Data presented one or pooled experiments (as indicated in individual figures). When pooled experiments are shown, all readings within the experiments were considered separately in the statistical analysis. Differences between means were compared using one- or two-way ANOVA followed by post-hoc pair-wise comparisons with Bonferroni or Dunnett adjustments as indicated (GraphPad Software, La Jolla, CA, USA). Significance was assessed relative to p < 0.05.

RESULTS

Validation of BRET assays and type I receptor homo-dimerization

In order to monitor interactions between the type I receptors of the TGFβ family, ALK1, ALK2, ALK3, ALK4 and ALK5 were fused at their C-termini to Renilla luciferase (Rluc) or the Aequorea victoria green fluorescent protein variant (GFP2). These receptors were chosen because they are ubiquitously distributed (ALK2, ALK4, ALK5) and probably co-expressed with other receptors [403, 404] or due to their known ability to form heterodimers with other type I receptors (ALK1, ALK2, ALK3, ALK5) [121, 131]. In the latter case we can look at those interactions as positive controls. On the other hand, ALK6 and ALK7 have a more restricted expression pattern [395, 405-408] and were not included in this preliminary assessment. Several of these constructs were tested using gene reporter assays and demonstrated to be functional before use as BRET donors or acceptors (Supplementary Figure S4.1A-E).

As proof of principle for the BRET assays, we carried out titration assays between ALK5-GFP2 (acceptor) and different Rluc tagged receptors as donors, including ALK5

(homodimer) and TGFBR2 (heterodimer). The β2-adrenergic receptor, β2AR-Rluc, and the T-cell co-receptor, CD8-Rluc, were used as presumptive negative controls, as these proteins are expressed at the plasma membrane but were not expected to interact with receptors of the TGFβ family. Cells were transfected with a fixed quantity of donor

158 (Rluc-expressing vectors) and increasing amounts of acceptor (ALK5-GFP2, in this case) to generate saturation curves.

BRET between ALK5-Rluc/ALK5-GFP2 or TGFBR2-Rluc/ALK5-GFP2 signals were saturable (Figure 4.1A), illustrating the specific homodimerization of ALK5 and its heterodimerization with TGFBR2. On the other hand, the signals obtained with CD8 and

β2AR did not saturate, suggesting that these interactions were non-specific, attributable to random associations between the receptors [409, 410]. In the particular case of β2AR, almost no BRET signal was observed when this receptor was used as a donor, despite the fact that BRET ratios were estimated using comparable luminescence values and these cells also expressed similar levels of acceptor (as evidenced by measuring the GFP2 expression, Figures 4.1B-C). However, a different outcome was observed when the β2AR served as acceptor molecule (see below).

Titration assays were also carried out after pre-treating the transfected cells with TGFβ1 or serum-free medium to assess whether the presence of ligand promoted changes in the BRET signal (Supplementary Figure S4.2). Contrary to what we expected, BRET saturation curves under basal and ligand-induced conditions were superimposable (i.e., there were no changes in the BRET50 or BRETmax values) for all the receptor combinations (Supplementary Figure S4.2A). These results suggest that, at least in the conditions used, TGFβ1 treatment did not foster receptor association or induce major conformational changes in the receptor intracellular domains [410-412].

Next we studied type I receptor homodimerization by BRET. Since it is known that type I receptors can interact with themselves, we used this information to assess whether the tagged receptors can form useful BRET pairs, i.e., determine if the orientation of the donor and acceptor moieties in these receptors allowed efficient resonance energy transfer. To assess type I receptor homodimerization, we transiently transfected HEK293 cells with a constant amount of the donor molecule (type I receptor tagged with Rluc) and increasing amounts of acceptor (type I receptor fused to GFP2). Again, CD8-Rluc was used as a negative control. All type I homodimers tested showed a higher BRET signal

159 compared to the negative control, indicating that they are potentially specific BRET pairs (Figure 4.2). However, the magnitude of BRET signals for different type I receptor homodimers was not equivalent, further suggesting that some acceptor-donor pairs are oriented more optimally for BRET; although this inconsistency might be partially explained by differences in the expression levels of the acceptor proteins (Supplementary Figure S4.3B, compare ALK3-GFP2 and ALK5-GFP2 fluorescence values).

Type I receptor heterodimerization

After verifying the ability of our constructs to homodimerize and generate BRET signals, we proceeded to evaluate the combinatorial possibilities of different type I receptor heterodimeric interactions. Here, we transfected HEK293 cells with different amounts of GFP2-tagged type I receptors in combination with a battery of donors, comprising the type I receptors ALK1 to ALK5 and CD8 as a negative control. We also included a set of conditions were β2AR-GFP10 served as acceptor to the different Rluc-labeled donors, as the β2AR proved to be a good negative control in our previous assays. We tried to obtain BRET ratios from samples exhibiting comparable levels of luminescence (RLU, Supplementary Figure S4.4) and fluorescence (RFU, Supplementary Figure S4.5) to prevent possible confounding effects due to variations in the protein expression levels. However, the expression of ALK3-GFP2 again was lower when compared to the other type I receptors, while β2AR-GFP10 was highly expressed.

Regardless of the disparity in fluorescence levels, all type I receptor combinations showed higher BRET values compared to the negative control (including those conditions where ALK3 was an acceptor), which indicate that these proteins have some intrinsic affinity for each other (Figure 4.3, summarized in Table 4.1). Nevertheless, certain BRET pairs showed higher signals than others and the energy transfer between a given BRET pair varied according to the identity of the donor and the acceptor. Overall, ALK1 behaved as a good acceptor, but when used as a donor, it showed reduced BRET with other receptors. ALK1 was able to dimerize with itself, as well as ALK2, 3, 4 and to a lesser extent ALK5 (Figure 4.3). ALK2 and ALK3 had similar combinatorial patterns,

160 showing stronger signals when associated with themselves (as homo- or heterodimers) or with ALK4. In addition, ALK2, but not ALK3, formed an effective BRET pair with ALK1 (Figure 4.3). On the other hand, although ALK4 appeared to be a promiscuous BRET donor, when used as an acceptor it showed higher BRET signals when associating with itself, followed by the interactions with ALK2 and ALK5. Likewise, in the experiments where ALK5 was the acceptor, it was able to interact with itself and ALK4, but no significant signal was detected with the other type I receptor tested, which suggests that it might be a poor BRET partner.

ALK4 and ALK5 interact with the β2-adrenergic receptor

When initially used as a donor, we failed to detect BRET between β2AR-Rluc and ALK5 (Figure 4.1). Ideally, in any BRET experiment, the tags should be alternated to confirm or refute particular interactions. Thus, we used β2AR as a BRET acceptor (β2AR-GFP10) when evaluating the heterodimeric interactions between the type I receptors. However, this time we noted all the type I receptors, but most strikingly ALK4 and ALK5, generated BRET in the presence of β2AR-GFP10 (Figure 4.3F). This experiment demonstrated the sensitivity of BRET to the position of the donor and acceptor moieties.

To resolve the issue of whether or not the β2AR could interact with type I receptors, we used myc-tagged receptors to compete BRET interactions between ALK5-GFP2 and ALK5-Rluc as an independent measure of their specificity. We observed that most type I receptors could efficiently compete the interaction (Supplementary Figure S4.6), confirming the promiscuous nature of type I receptor associations. Interestingly, at higher levels of expression, myc-tagged β2AR also competed the BRET interaction, implying that β2AR can indeed interact with ALK5 and this BRET pair only functions when donor and acceptor moiety are positioned in one particular orientation.

In order to corroborate the potential physical association between β2AR and ALK5, we determined whether these receptors could be co-precipitated when expressed in HEK293 cells. In this case, we used HEK293 cells stably expressing β2AR receptor fused to a streptavidin-binding protein and HA-motif (HEK-TAP-β2AR), and transiently transfected

161 them with the ALK5-myc construct. We found that purification of β2AR on streptavidin beads also pulled down ALK5-myc (Figure 4.4A). Likewise, β2AR was co- immunoprecipitated with ALK5, validating the interaction between the two proteins

(Figure 4.4B). Interestingly, association of β2AR with type I receptors seemed to be specific for ALK4 and ALK5, as we failed to strongly co-precipitate other type I receptors with β2AR (Figure 4.4C). These data were consistent with our BRET results. Taken together, these data suggest that receptors of the TGFβ superfamily (specifically

ALK4 and ALK5) can associate with a G protein-coupled receptor, β2AR.

DISCUSSION

Using BRET, we showed that ALK4 has the potential to form heterodimers with ALK2 and ALK5. Moreover, we provide evidence indicating that β2AR physically interacts with the type I receptors ALK4 and ALK5. BRET methodology have been successfully employed to study oligomerization of GPCRs [401, 413, 414] and dimerization of specific TGFβ receptors pairs [137, 140]; yet, to our knowledge this report constitutes the first systematic assessment of possible type I receptor combinatorial interactions.

The results presented here show that the type I receptors studied are able to oligomerize with each other, suggesting that there are no major structural restrictions that limit their association. However, certain BRET pairs yielded higher signals than others. The discrepancy in the magnitude of the BRET signal could be caused by reduced resonance energy transfer efficiency or differences in the affinity of receptor-receptor interactions. Unfortunately, these particular BRET experiments, as designed, do not yield information regarding the affinity of the receptor associations [415].

We detected type I receptor heterodimers already reported in the literature, such as ALK2-ALK3 [137]; but, we had difficulty confirming ALK1-ALK5 dimerization, which has been described in both chondrocytes [139, 416] and endothelial cells [121, 138]. ALK1 and ALK5 dimerize in the absence of ligand but, at least in endothelial cells, the interaction is dependent upon the co-expression of a co-receptor, endoglin [121, 139]. As

162 endoglin levels in HEK293 are low or absent [417, 418], we examined the effects of endoglin co-expression on ALK1-ALK5 dimerization; however, this failed to modify the BRET signal (data not shown). These data suggest that the relative orientation of the acceptor and donor moieties when fused to ALK1 and ALK5 might not be favourable for BRET. Similarly, ALK5-ALK2 and ALK5-ALK3 dimers generated only weak BRET signals in our assays, although these pairs have been reported in different cell lineages, including HEK293 cells [131]. It is possible that these combination are not permissible in all cell types (even different HEK293 subclones) and/or specific receptor-interacting proteins are required to allow ALK5 interaction with ALK2 and ALK3, as it has been previously suggested [131]. Alternative placement of the tags might help resolve these issues.

When conducting the BRET saturation experiments in the presence of TGFβ1, we did not detect ligand-driven changes in the association of ALK5 with TGFBR2 [22, 419] or the conformational changes reported in the ALK5 intracellular domain upon ligand binding [20, 21]. Possibly a large fraction of the overexpressed receptors were already associated in pre-formed complexes [419, 420] or ligand-driven receptor association were lost during sample preparation. Given that the substrate for Renilla luciferase is cell- permeable, another possibility is that BRET signals came mainly from receptor complexes located in intracellular compartments, which cannot respond appropriately to ligand induction [421]. Experiments using broken membrane preparations might allow us to test this directly. Additionally, the Rluc and GFP2 tags might again be inappropriately positioned to detect conformational changes in the receptors. In any case, it seems that although BRET allows us to detect the association between receptors in the TGFβ family, our experimental approach has certain limitations that prevent tracking dynamic changes in receptor interactions. This will certainly need to be optimized in future experiments, probably by conducting real-time BRET assays [409, 422] or by using alternative locations for introducing BRET donor and acceptor moieties [423, 424].

Interestingly, we observed novel receptor combinations, such as ALK2-ALK4 and ALK4-ALK5. However, saturation and competition assays in conjunction with other

163 techniques, such as co-immunoprecipitation, should be performed to validate these possible receptor interactions. Of course, the signaling consequences should also be evaluated for these new combinations.

ALK2 and ALK4 are distant phylogenetically, [29]. ALK2 can bind activins [13, 16, 425], but does not appear to propagate their signals [14]. In fact, overexpression of ALK2 leads to a reduction in activin signaling mediated by SMAD2/3 activation in different systems [261, 426]. Therefore, it would be expected that ALK2 might function as a dominant negative receptor when associated with ALK4. Alternatively, activin binding to ALK2-ALK4 complexes might trigger alternative signaling pathways, as in the case of ALK2-ALK5 heterodimers [131].

ALK4 and ALK5, on the other hand, bind GDF-8 (myostatin) [427] and GDF11 [428]. In addition, these receptors are ubiquitously distributed [403, 404] and they exhibit high sequence homology, so it is reasonable to think that they might oligomerize. Given that both receptors activate SMAD2/3, ALK4-ALK5 dimers would probably elicit similar responses relative to ALK4 or ALK5 homodimers. However, differences in ALK4 and ALK5 regulation [for instance, ALK5 can be sumoylated while ALK4 cannot [82]], might contribute to generate receptor complexes with unique signaling characteristics.

We also found converging evidence (BRET, affinity pull-down, immunoprecipitation) that supports the existence of ALK4/β2AR and ALK5/β2AR complexes. Furthermore, protein fragment complementation assays, based on reconstitution of Venus fluorescent protein, indicate that β2AR-β2AR and ALK5-β2AR complexes locate at the plasma membrane and some intracellular vesicles (Supplementary Figure S4.7B-C). Interestingly, when ALK5 split Venus constructs were expressed alone the fluorescent signal was mostly found at the nuclear/perinuclear region (Supplementary Figure S4.7A), which might represent an artifact due to overexpression or the nuclear accumulation of ALK5 intracellular domain upon receptor cleavage [109].

164 The potential interaction between ALK4/5 and β2AR is particularly interesting because those receptors colocalize in different tissues such as heart and the pituitary [429-432] and there is evidence of crosstalk between TGFβ- and β2AR-mediated signaling, although the effects of these interactions seems to be cell-dependent [210, 433, 434]. It is possible that the signaling crosstalk occurs at the level of the effectors or that association of TGFβ receptors with β2AR alters their trafficking properties. In this regard, it has been reported that, upon parathyroid hormone (PTH) stimulation, the PTH receptor (another GCPR) interacts with TGFBR2 and the two receptors are co-internalized, leading to attenuation of TGFβ and PTH signaling [207]. The functional significance of these interactions has yet to be addressed (see Chapter 5).

Although we acknowledge the caveat that the information derived here relies on overexpressed proteins, a situation that might not be physiologically relevant, these observations provide hints about the extent of combinatorial possibilities in the TGFβ superfamily and may help us discern if targeting a particular receptor may elicit unexpected secondary effects. Furthermore, our data shows that some family members can interact (either directly or indirectly) with β2AR, adding further complexity to both TGFβ and GPCR signaling. It remains to be determined if the receptor dimers proposed here are functional and in which circumstances they would be formed.

165 Acknowlegments

This work was supported by grants from the Canadian Institutes of Health Research (CIHR) to T.E.H (MOP-36279 and MOP-79354), and D.J.B (MOP-89991). T.E.H. is a Chercheur National and D.J.B. a Chercheur Boursier Senior of the Fonds de la Recherche en Santé du Québec (FRSQ). C.R. was supported by a scholarship from the McGill CIHR Drug Development Training Program.

166 Figure legends

Figure 4.1. Saturation curve for ALK5 homo and heterodimerization. (A) HEK293 cells were transfected with a fixed amount of Rluc expressing vectors (0.1 or 0.2 μg) and different amounts of ALK5-GFP2 (0; 0.125; 0.25; 0.5; 1 or 2 μg). CD8-Rluc (single pass transmembrane receptor) and β2AR-Rluc (a G protein-coupled receptor) were used as negative controls. BRET background was set to the conditions where Rluc tagged proteins were expressed alone and those values were subtracted to yield net BRET. Each point represents the mean +/- SEM, n = 3. The curves were fitted using non-linear (ALK5, TGFBR2) or linear (CD8, β2AR) regression (GraphPad Prism). (B) Relative luminescence values (RLU) shown demonstrates that expression levels of the donors were constant. (C) Relative fluorescence values (RFU) demonstrate that expression levels of the donors increase at similar rates under the different conditions.

Figure 4.2. TGFβ type I receptor homodimerization. HEK293 cells were transfected with 0.5 μg of Rluc expressing vectors and three different concentrations (0.5, 1 or 1.5 μg) of the corresponding type I receptor fused to GFP2. CD8-Rluc was utilized as negative control. (A) ALK1-GFP2; (B) ALK2-GFP2; (C) ALK3-GFP2; (D) ALK4-GFP2 and (E) ALK5-GFP2. BRET ratios were calculated by dividing amount of light emitted through the 515/35 filter over that passed by the 410/80 nm. Shown are mean +/- SEM of three independent experiments with three readings per experiment (n = 9). Dunnett multiple comparison test ($ p < 0.001) versus CD8-Rluc control.

Figure 4.3. TGFβ type I receptor heterodimerization. HEK293 cells were transfected with 0.5 μg of Rluc-expressing vectors and three different concentrations (0.5, 1 or 1.5

μg) of the corresponding type I receptor tagged to GFP2 or β2AR-GFP10. CD8-Rluc was utilized as negative control. (A) ALK1-GFP2; (B) ALK2-GFP2; (C) ALK3-GFP2; (D)

ALK4-GFP2; (E) ALK5-GFP2 and (F) β2AR-GFP10. BRET ratios were calculated by dividing amount of light emitted through the 515/35 filter over that passed by the 410/80 nm. Shown are mean +/- SEM of 3-5 independent experiments with three readings per

167 experiment (n = 9-15). Dunnett multiple comparison test (* p < 0.05; # p < 0.01; $ p < 0.001) versus CD8-Rluc control.

Figure 4.4. Co-immunoprecipitation of β2AR and ALK5 in HEK293 cells. HEK293 cells stably expressing a tagged version of the β2-adrenergic receptor (HEK-TAP-β2AR) were transfected with ALK5-myc (A and B), or the indicated myc-tagged type I receptors (C). Forty-eight hours after transfection crude membrane preparations were obtained and used for streptavidin bead purification (A and C) or anti-myc immunoprecipitation (B), as described in MATERIALS AND METHODS. Whole cell lysates were analyzed by immunoblot using the indicated antibodies. Images shown are from a representative experiment.

168 Supplementary Figure legends

Supplementary Figure S4.1. Testing the functionality of tagged type I receptors. Gene reporter assays. (A-B) LβT2 cells seeded in 24-well plates were transfected with murine -1990/+1 Fshb-luc reporter (450 ng/well) along with 100 ng of the indicated receptor expression vectors. One day post-transfection, cells were placed in serum free medium (A) or treated with 25 ng/ml activin A (B), for 24 hours. Treatments were performed in triplicate. Results are from single experiments and are presented as the fold- stimulation by BMPR2 over-expression (A) or activin A treatment (B) in the presence of each type I receptor (C) L17 cells grown in 24-well plates were transfected with 450 ng/well CAGA-luc and 50 ng/well ALK5-myc. The next day, cells were treated with 5 ng/ml TGFβ1 in serum free medium for 24 hours. Treatments were made in triplicate. Data represent mean (+/-SEM) of three independent experiments. (D) Same as (C) but cell were transfected with 450 ng/well 3TP-luc and 100 ng/well ALK5-Rluc. Treatments were performed in duplicate. Data represent mean (+/-SEM) of two independent experiments. (E) DR26 cells cultured in 24 well plates were transfected with 450 ng/well 3TP-luc along with 100 ng/well TGFBR2-Rluc. Cells were treated the next day with 5 ng/ml TGFβ1 in serum free medium for 24 hours. Treatments were performed in duplicate. Results are from a single experiment. The data from (C-E) are presented as fold-stimulation by TGFβ1 in the presence of each receptor.

Supplementary Figure S4.2. Effect of TGFβ1 treatment on ALK5 homo and hetero- dimerization. (A) HEK293 cells were transfected with 0.2 μg (ALK5, TGFBR2, CD8) or 0.1 μg (β2AR) of Rluc expressing vectors and different amounts of ALK5-GFP2 (0; 0.125; 0.25; 0.5; 1 or 2 μg). To assess the effect of the ligand, cells were incubated with serum-free media (SFM) or 5 ng/mL TGFβ1 in SFM for 20 minutes before conducting the BRET assay. BRET background was set to the conditions where Rluc tagged proteins were expressed alone and those values were subtracted to yield net BRET. Each point represents the mean +/- SEM of two independent experiments, each one with three readings per sample (n = 6). The curves were fit using non-linear (ALK5, TGFBR2) or linear (CD8) regression (GraphPad Prism). The estimated BRETmax and BRET50 values

169 (+/- SD) are summarized in the accompanying Table. (B) Relative luminescence values (RLU) shown demonstrate that expression levels of the donors were constant. (C) Relative fluorescence values (RFU) demonstrate that expression levels of the donors increase at similar rates under the different conditions.

Supplementary Figure S4.3. Relative luminescence (A) and fluorescence (B) values corresponding to the cells used in Figure 4.2. The relative luminescence values (RLU, A) shown illustrate that expression levels of the donors were constant and relative fluorescence values (RFU, B) demonstrate that expression levels of the donors increase at similar but not identical rates under the different conditions, except for ALK3-GFP2.

Supplementary Figure S4.4. Relative luminescence values (RLU) corresponding to the cells used in Figure 4.3. The relative luminescence values shown demonstrate that expression levels of the donors were comparable under the different conditions. (A) ALK1-GFP2; (B) ALK2-GFP2; (C) ALK3-GFP2; (D) ALK4-GFP2; (E) ALK5-GFP2 and (F) β2AR-GFP10.

Supplementary Figure S4.5. Relative fluorescence values (RFU) corresponding to the cells used in Figure 4.3. The relative fluorescence shown values indicate that expression levels of the donors increase at similar rates under the different conditions. (A) ALK1-GFP2; (B) ALK2-GFP2; (C) ALK3-GFP2; (D) ALK4-GFP2; (E) ALK5-

GFP2 and (F) β2AR-GFP10.

Supplementary Figure S4.6. Competition of the ALK5 homodimer in the presence of different type I receptors and β2AR. HEK293 cells were transfected with 0.4 μg of ALK5-Rluc, 1 μg of ALK5-GFP2 along with two different concentrations (0.75, 1.5 or

3.0 μg) of different myc-tagged type I receptor or β2AR-myc-expressing vectors. In the control condition, cells were co-transfected with 3.0 μg of pcDNA3.1. (A) BRET ratios were calculated by dividing amount of light emitted through the 515/35 filter over that passed by the 410/80 nm. Shown are mean +/- SEM of three readings coming for one

170 experiment. (B) and (C) are the corresponding luminescence (B) and fluorescence (C) values for the cells used.

Supplementary Figure S4.7. β2AR colocalize with ALK5 at the plasma membrane. HEK293 seeded in 6 well plates were co-transfected with the following pairs: (A) ALK5-

Venus1 and ALK5-Venus2, (B) β2AR-Venus1 and β2AR-Venus2 or (C) β2AR-Venus1 and ALK5-Venus2 constructs using a conventional calcium phosphate co-precipitation protocol. Forty eight hours after transfection reconstitution of Venus was assessed by confocal microscopy. Each figure shows three representative cells from a single experiment.

171 Figure 4.1

A

B

C

172 Figure 4.2 A B

C D

E

173 Figure 4.3

A D

B E

C F

174 Figure 4.4

A

B

C

175 Supplementary Figure S4.1 A B

C D E

176 Supplementary Figure S4.2

A

ALK5-ALK5 ALK5-ALK5 ALK5-TGFBR2 ALK5-TGFBR2

(No ligand) (TGFβ1) (No ligand) (TGFβ1)

BRET50 4.48 ± 0.44 4.98 ± 0.27 3.47 ± 0.54 4.32 ± 0.52

BRETmax 0.62 ± 0.02 0.68 ± 0. 01 0.20 ± 0.01 0.20 ± 0.01

B

C

177 Supplementary Figure S4.3

A B

178 Supplementary Figure S4.4 A B C

D E F

179 Supplementary Figure S4.5 A B C

D E F

180 Supplementary Figure S4.6

A

B

C

181 Supplementary Figure S4.7

A

B

C

182 Table 4.1: Summary of type I receptor interactions as evaluated by BRET. Donor/Acceptor ALK1 ALK2 ALK3 ALK4 ALK5 β2AR ALK1 ++ + + + - + ALK2 ++ ++ ++ +± - + ALK3 ++ + +± + - + ALK4 ++ ++ ++ +++ + ++ ALK5 + + + +± +++ ++

The signs represent a qualitative representation of the BRET results obtained in Figures 2 and 3.

183 Chapter 5: General discussion

Activins are members of the TGFβ family originally identified for their role in FSH expression and secretion [246, 247]. Since then, the list of activin functions has tremendously expanded. These functions are cell context- and dose-dependent, and they include: regulation of stem cell proliferation [435, 436], pancreatic β-cell proliferation and function [437, 438], erythropoiesis [439], ovarian follicular development [440, 441], bone formation [442], control of immune response [443, 444], and neuroprotection [445], among others. As a result of these pleiotropic effects, activins have also been associated with numerous pathological conditions and there is a growing interest in the characterization of activin signaling pathways and their modulation for therapeutic purposes [220, 221].

As a member of the TGFβ superfamily, activin signaling is tightly controlled, again in a context-specific manner. For instance, the antagonist FST and inhibins prevent activin- receptor interactions [49, 116, 264] and expression of ARIPs and ubiquitin ligases modulate receptor levels at the plasma membrane [89, 102, 165]. Additionally, activin pathways and other TGFβ signaling cascades are modulated at the intracellular level by many factors including the I-SMADs [165, 333] and interaction with cell-specific transcriptional factors (reviewed in [195]). However, despite our vast knowledge about activins signaling and physiological relevance, novel regulatory systems continue to be reported.

Here, we have described additional mechanisms whereby activin function could be modulated in gonadotropes. One of them involves the rapid regulation of ACVR2 protein levels, presumably by receptor shedding at the plasma membrane (Chapter 2); the other a novel interaction between activin and BMPR2, a type II receptor not initially associated with activin signaling (Chapter 3). Moreover we showed preliminary evidence of ALK4 heterodimerization with ALK2 and ALK5, as well as the physical association between

ALK4 and ALK5 with the β2AR (Chapter 4).

184 5.1 ACVR2 turnover in LβT2 cells

In Chapter 2, we estimated that the half-life of ACVR2 in LβT2 cells was approximately 2 hours. Due to rapid ACVR2 turnover, in the absence of protein synthesis, activin signaling is quickly impaired leading to decrease in Fshb expression, which explains the inhibin-like effect of CHX in gonadotropes. We also found, contrary to what has been reported for other receptors in the family, that mature ACVR2 is not degraded via either the lysosome or proteasome in LβT2 cells. Instead, our preliminary data suggested that membrane shedding is a key determinant in ACVR2 degradation, based on the detection of protein fragments with similar sizes to ACVR2 intracellular and extracellular domains in cell lysates and media, respectively. The identity of those fragments could be determined by excising the corresponding bands from acrylamide gels and evaluating the samples using N-terminal sequencing (Edman degradation) and mass spectrometry. This information will help us to confirm the putative shedding of ACVR2 and identify the sheddase’s cleavage site(s). Provided that ACVR2 is shed, we could conduct similar experiments in other cell types to determine if this represents a cell-specific phenomenon as well as ascertain the necessary and sufficient components in the process.

We need to further characterize ACVR2 shedding and identify the enzyme(s) responsible for receptor cleavage. Shedding is usually a regulated process and it can be induced by several mechanisms, PKC activation by phorbol esters (like PMA) is perhaps the best characterized of these mechanisms [446, 447]. Preliminary experiments suggest that ACVR2 levels are not affected by PMA treatment (data not shown) in LβT2 cells. Likewise, ectodomain shedding of the TGFβ co-receptor, betaglycan, is PMA-insensitive, but rather induced by treatment with the tyrosine phosphatase inhibitor, pervanadate [375, 448]. Therefore, we need to test the effects of diverse stimuli, associated with shedding activation, including pervanadate, calcium ionophores or cytokines (IL-1β; [446, 447] to further understand how ACVR2 ectodomain cleavage is regulated.

The majority of the sheddases that mediate hydrolysis of cell surface proteins are zinc- dependent metalloproteases from the disintegrin and metalloproteinase (ADAM) family

185 [449, 450], although members of the matrix metalloproteinase family can play a role as well [375, 446, 447]. In humans, 21 ADAM genes have been identified, but only 12 of these code for proteins with a typical Zn-binding active site and are presumed to possess functional proteolytic capacity [449]. Members of the ADAM family contain a highly conserved catalytic domain, they are able to process a wide range of substrates (including ALK5) and most are inhibited by hydroxamic acid-based compounds, such as GM6001, BB94 and TAPI [450, 451]. Our results suggest that ACVR2 degradation is refractory to GM6001 treatment (Chapter 2). It is possible that ACVR2 ectodomain hydrolysis is mediated by a GM6001-insensitive proteinase, similar to one of the glycoprotein VI sheddases [452]. Alternatively, ACVR2 hydrolysis might be controlled by an aspartate protease, such as BACE1.

BACE1 is an integral membrane glycoprotein, originally identified as the β-secretase responsible of amyloid precursor protein cleavage and production of a 42-residue amyloid peptide (Aβ42) in Alzheimer’s disease [453]. BACE1 is more restrictive than the ADAMs in terms of substrate selectivity and among its targets are: neuronal cell adhesion molecule 1, toll-like receptor 9, neogenin, the -like growth factor 2 receptor and interestingly, the pseudo-receptor, BAMBI [454]. Nevertheless, β-secretase processing has not been reported for any TGFβ receptor. Since metalloproteinases and β-secretases have different pharmacological profiles, treatment of LβT2 cells with diverse inhibitors in conjunction with information yielded by N-terminal sequencing of the proteolytic products will facilitate the identification of the presumptive ACVR2 sheddase.

BMPR2, which we identified as a novel activin type II receptor in Chapter 3, seems to also be turned over rapidly in gonadotropes. Both endogenous BMPR2-L and transiently transfected BMPR2-S tagged with GFP2, are barely detectable after 6 hours of CHX treatment (Fig. 5.1). It has been reported that BMPR2 can be degraded via the proteasome or lysosome [98, 371]. In addition, BMPR2 is shed following treatment with PMA [110] in primary human bone cells. Therefore, if ACVR2 is cleaved at the plasma membrane by a process independent of PKC activation, cells could potentially alter expression of BMPR2 without affecting ACVR2 levels and vice versa.

186

Shedding would restrict activin signaling both by decreasing the amount of available intact receptor at the plasma membrane and, in the case of ACVR2, by releasing the ECD, which may bind and bioneutralize the ligand [377]. In cells that are responsive to activin, as is the case of hepatocytes, receptor shedding might function as a protective mechanism against excessive activin signaling. For instance, liver inflammatory and fibrotic processes lead to an increase in autocrine/paracrine activin synthesis, which if disproportionate, could affect hepatocyte growth and impair liver regeneration [455, 456]. Additionally, receptor shedding might also contribute with the establishment and control of activin/nodal/BMP morphogen gradients during embryogenesis. On the other hand, alterations in the expression of activin receptors and activin responses have been observed in pathological conditions such as cancer and in the dysfunctional human testis [457, 458]. Therefore understanding the precise mechanisms underlying ACVR2 degradation and shedding could yield an important therapeutic target for these and other pathologies. For instance, inhibitors of ACVR2 sheddase could be used in assisted reproduction, to increase endogenous activin signaling at the level of the pituitary and in turn, FSH production. Conversely, activators of ACVR2 shedding, could be used instead of the ACVR2-IgG-Fc fusion protein for treatment of anemia and bone loss [459], or to prevent bone metastasis in patients with breast cancer [460]

5.2 BMPR2 as activin receptor in LβT2 cells

Data from Chapter 3 showed that BMPR2 is a bona fide activin receptor in gonadotropes. The BMPR2-activin interaction seems to be specific since activin did not bind either TGFBR2 or AMHR2 (Fig. 3.1C) and at the same time the BMPR2-ECD partially antagonized activin A-, but not TGFβ1-induced effects on CAGA luciferase reporter activity (Fig. 3.4).

According to our results, only the BMPR2 short isoform (BMPR2-S) is able to propagate activin signals to regulate Fshb promoter activity. However, we did not evaluate the possibility that activin binding to BMPR2 (short or long isoform) could lead to activation

187 of SMAD-independent signaling pathways. The BMPR2 long isoform differs from the short one by the presence of a long C-terminal tail following the kinase domain [9, 10, 461]. Both isoforms are able to interact with type I receptors and have shown to be active in diverse functional assays [9, 10, 462, 463]. The long C-tail present in BMPR2-L facilitates its interaction with diverse proteins, such as Trb3 [464], LIM kinase 1 [70] and cGMP-dependent kinase 1 [69] which modulate canonical BMP signaling or facilitate crosstalk with other signaling pathways. In addition, GST pulldown followed by mass spectrometry showed that BMPR2-S and BMPR2-L co-precipitate distinct subsets of interacting proteins [71], indicating that BMPR2 isoforms have distinct signaling capacities.

In our knockdown experiments, we noted that targeting Bmpr2 expression with two different siRNAs (#1 and #4) impaired activin-induced Fshb and CAGA-luc reporter activities (Supplementary Figure S3.6). In contrast, a third siRNA (#2) had no effect in those systems. Based on their target sequences, we presume that siRNA #1 and #4 target both murine BMPR2 isoforms (notice that in our assays we used a human-BMPR2-S expression construct), while siRNA #2 affects only BMPR2-L expression. However, we ignore if BMPR2-S is expressed in murine gonadotropes. Northern blot analysis indicates the presence of up to three different BMPR2 transcripts in various human and murine tissues [9, 391, 465], suggesting that BMPR2 isoforms may be widely expressed. We will use northern blots with specific probes that anneal towards the 5’ or 3’ end of BMPR2-L to clearly establish whether or not BMPR-S is expressed in LβT2 and pituitary cells. In addition, rapid amplification of cDNA ends (RACE) using RNA samples isolated from LβT2 cells or purified gonadotropes, along with BMPR2 specific primers, could provide the sequence of BMPR2 isoforms.

We would like to conduct similar experiments to the ones described in Chapter 3, using different cell types, such as granulosa cells, to determine if BMPR2 can also transduce activin signals in these systems and if so, what factors might be required for the interaction to take place. It is possible, for example, that the presence of accessory proteins favors activin binding and signaling through BMPR2 in an analogous manner as

188 RGM proteins enhance ACVR2 utilization by BMP2/4 [127] or Cripto facilitates GDF8 siganling via ALK4 in myogenic cells [427].

Another question that should be addressed is the putative, yet plausible interaction between ALK4 and BMPR2. We observed that in the presence of the ALK4/5/7 inhibitor, SB421543, BMPR2-S over-expression did not potentiate activin A-induced Fshb expression (Chapter 3). Given that ALK4 is the only activin A receptor reported to date [261], these results suggest that BMPR2 effects on activin signaling requires the presence of this receptor. One way to confirm if ALK4 dimerizes with BMPR2 would be to assess Fshb reporter activity after activin A treatment, in LβT2 cells transiently transfected with BMPR2-S along and Acvr1b (Alk4) siRNA. Alternatively, a more direct approach to confirm ALK4-BMPR2 association would be using 125I-activin A to affinity label CHO cells transfected with BMPR2 and/or ALK4 expression vectors, and evaluate the presence of ALK4-BMPR2 complexes by SDS PAGE/autoradiography.

To determine the role of BMPR2 in mediating activin regulated FSH synthesis in vivo, it would be necessary to develop a gonadotrope-specific Bmpr2 knockout mice model, by crossing the Gnrhr-Cre (GRIC) [272] and the Bmpr23loxP/3loxP [466] transgenic mice, since the Bmpr2 global knockout is embryonic lethal [467]. These animals would express a mutant BMPR2 in gonadotropes, which lacks the transmembrane domain and part of the kinase domain, thus affecting BMPR2-S and -L expression [466]. Once efficient Cre/lox recombination in gonadotropes is confirmed by qPCR, measurement of circulating FSH levels and Fshb mRNA expression in pituitary should be conducted to discern the potential role of BMPR2 in Fshb transcriptional regulation. According to the results presented in this dissertation, and in previous experiments performed in our lab [325], we would expect a reduction in Fshb mRNA levels after Bmpr2 ablation, since this receptor can mediate both BMP2 and activin stimulatory signals in gonadotropes. However, given that BMPR2-L is expressed in gonadotropes and this isoform can bind activins but not propagate their signals (at least in the context of Fhsb transcription), knocking out Bmpr2 might, in the end, lead to an increase in activin availability and signaling through ACVR2. In addition, disruption of Bmpr2 expression will differentially affect signaling

189 of various BMP ligands, which induce mFshb transcription with different potency [282, 468]. Therefore, the weighted contribution of those different components would determine if Fshb transcription decreases, increases or remains unaffected.

5.3 BMPR2 ligand binding surface

Based on the crystal structure of activin and ACVR2B, we generated a hypothetical model of activin A bound to BMPR2 (Figure 3.2). According to this model, BMPR2 makes similar ligand binding contacts with activin A as it does to BMP2 [63]. BMPR2 and ACVR2B interactions with activin A seems to be largely mediated by a hydrophobic surface called loop A, although the loop A from ACVR2B is five amino-acids shorter [40, 63].

In order to further validate this model, we generated a mutant receptor called BMPR2- swap, in which we substituted five residues from the BMPR2-ECD domain (located in the A-loop), with the corresponding residues from ACVR2B ECD domain (Fig. 5.2A).

BMPR2-swap was expressed at similar levels as BMPR2-S-WT (Fig. 5.2B). In LβT2 cells, BMPR2-swap actually conferred heightened activin-dependent and -independent Fshb reporter promoter activity (Fig. 5.2C), similar to what we observed in cells transfected with ACVR2B (Supplementary Fig. S3.3A). These data suggest that not only is there structural commonality between BMPR2 and ACVR2B ligand binding interfaces, but also that only a small structural motif (five to eight residues) significantly affected the ability of BMPR2 to transduce activin signals, yielding a more ACVR2B like-receptor. It remains to be established if swap mutation has an impact on activin binding to BMPR2.

5.4 Type I receptor interactions

Heterodimerization of TGFβ receptors can generate signaling diversity. The best example studied to date has been ALK5-ALK1 dimerization in endothelial cells and chondrocytes, which leads to phosphorylation of SMAD1/5 in response to TGFβ treatment, in addition

190 to canonical induction of SMAD2/3 phosphorylation [121, 138, 139]. Also, oligomerization of the BMP type I receptor ALK3 with ALK2 and ALK6 has been reported [130, 137], even though the functional consequences of these interactions were not examined. Less is known about oligomerization of activin type I receptors. In Chapter 4, our BRET results suggest that ALK4 can oligomerize with ALK2 and ALK5 in transiently transfected HEK293 cells; however, these putative ALK4 heterodimers should be validated using complementary methods.

Co-immunoprecipitation assays, using receptors labeled with different tags, could be performed to confirm ALK4 physical interaction with ALK2 or ALK5. Because dimerization might be induced or altered (in terms of stability) by the presence of ligand [130] these co-immunoprecipitation assays should initially be conducted in the presence of activin A or GDF8, depending if ALK2-ALK4 or ALK4-ALK5 dimers, respectively, are being studied, [13, 16, 425, 469].

Binding assays conducted in cells transfected with each receptor alone or in combination, followed by crosslinking and immunoprecipitation assays would indicate if these receptor complexes localize to the plasma membrane. In this case, cells should be transfected with an appropriate type II receptor, such as ACVR2B, because neither ALK4, nor ALK2 or ALK5 can bind ligand by themselves [14, 16, 425, 469]. Additionally, subcellular fractionation of cell lysates using sucrose density gradients and analysis of these samples by immunoprecipitation and immunoblot (under reducing and non-reducing conditions) would help to determine where and when these receptor complexes are formed [18, 130].

Another dimer that would be interesting to study, but was not covered in our study, is ALK4-ALK7. ALK4 is ubiquitously expressed [403], whereas ALK7 is mostly found in brain [405, 406]. Interestingly, both receptors are expressed in pituitary [261, 299]. ALK4 propagates signals from all activins (A, B and AB), while ALK7 only transduces activin B and AB signals [260, 261]. It has been reported that expression of ALK7 actually increases activin -AB and -B responses in HEK293, HT22, and LβT2 cells, which already express ALK4 [260, 261, 266]. Since ALK7 is unable to bind activin A (INHBA

191 homodimer) [406] and activin AB is a heterodimer of INHBA and IBHBB subunits, it is puzzling how ALK7 could be more efficient than ALK4 in propagating activin AB signals. Moreover, potentiation of activin AB signaling by ALK7 expression is not completely reversed by the presence of a dominant negative form of the receptor [260]. Hence, it is possible that activin AB signals via ALK4-ALK7 dimers, in addition to ALK4 and ALK7 homodimers. Alternatively, activin A dimerization with activin B could cause a conformational change that fosters interaction with ALK7, as suggested elsewhere [261].

The functional implications of ALK4-ALK7 dimerization could be evaluated using the same approach described by Lavery et al. [137] to validate ALK3-ALK6 dimerization. In this case, we should use a cell line that expresses both receptors and measure an activin AB or activin B-induced response (such as induction of CAGA-luc reporter activity) after knocking down ALK4, ALK7 or both receptors simultaneously using targeted siRNA. Once receptor knockdown is confirmed by qPCR, we could calculate if the inhibition of activin-stimulated responses caused by decrease in ALK4 and ALK7 expression is additive or if there is a fraction of the response that requires the expression of both receptors. These experiments could be conducted in LβT2 cells [261, 299], but they are notoriously difficult to transfect [305].

Given that Acvr1c (ALK7) knockout mice do not show any fertility impairment, the potential role of ALK7 in mediating activin B-regulated FSH synthesis in vivo seems negligible, although it is important to note that circulating levels of FSH protein were not determined in these animals [303]. ALK4-ALK7 dimers could play a role in regulation of pancreatic β-cell function. In these cells, activin A and B have antagonizing effects on glucose-dependent Ca2+ influx, which may be mediated by different type I receptor complexes [470].

A question that goes beyond the scope of this thesis but would be interesting to answer is where receptor dimerization takes place (ER, plasma membrane, both). If dimerization of type I receptors occurs early, at the ER, then structural constraints and distinct tissue

192 distribution would primarily limit formation of certain receptor combinations. However, if ligand binding is the key element triggering receptor dimerization, then not only the receptor structures will play a role, but also the type and concentration of ligands present in the milieu and which receptors can they recruit. In any case, although it is still not clear which domains mediate interactions between receptors, the factors that determine receptor combination specificity likely involve all of the mechanisms mentioned above. Our results from Chapter 4 suggest that the structural features of type I receptor are not the primary determinants of receptor association and the presence of cell-specific factors might be critical regulators in this process.

5.5 β2AR interaction with TGFβ receptors

β2AR and ALK4/5 are co-expressed in the heart [471], where they play different roles in myocardium contractility [472, 473] and remodelling [474, 475]. There is evidence of crosstalk between TGFβ and β-adrenergic signaling pathways. For instance, in transgenic mice over-expressing TGFβ1, there is an increase in expression of β-adrenergic receptors in the myocardium accompanied by cardiac hypertrophy [433]. However, TGFβ1 treatment reduces β2AR cell surface expression in cultured human tracheal smooth muscle cells [210] and rat alveolar epithelial cells [434]. Although the mechanisms underlying these responses have not been completely elucidated, signaling crosstalk between TGFβ and adrenergic pathways can take place at different levels, and its effects are cell-specific.

In Chapter 4 we reported the association between the β2AR and the TGFβ type I receptors ALK4 and ALK5 using BRET analysis and co-precipitation. These interactions seem to be specific, since ALK1 and ALK3 showed no sign of physical interaction with

β2AR. We also showed by protein fragment complementation assays, that ALK5 and

β2AR oligomers locate at the plasma membrane.

In order to study the functional significance of ALK4/5-β2AR interactions, we transiently transfected HEK293 cells with a β2AR expressing construct and evaluated levels of

193 SMAD2 phosphorylation induced by activin or TGFβ in the presence or absence of the βAR agonist, isoproterenol (Fig. 5.3). After 30 minutes of treatment, isoproterenol did not alter levels of ligand-stimulated SMAD2 phosphorylation, indicating that β2AR activation has no apparent immediate effects on canonical activin/TGFβ signaling. However, we can not discount the possibility that in this timeframe β-adrenergic stimulation affects a TGFβ/activin SMAD-independent pathway.

Curiously, incubation of HEK293 cells with isoproterenol for 24 hours did not affect activin A- or TGFβ1-induced CAGA-luc reporter activity, although the fold induction was reduced by virtue of a small increase in basal activity (Fig. 5.4). This result might be due to low levels of endogenous β2AR expressed in HEK293 cells. β2AR overexpression, on the other hand, dramatically reduced activin- and TGFβ-induced CAGA-luc reporter activity regardless of isoproterenol treatment (Fig. 5.4). Taken together, these results suggest that if β2AR interaction has any effect on ALK4/ALK5 functionality, it does not require ligand activation. Another possibility is that β2AR overexpression actually lead to ligand-independent activation of the receptor, which has been previously reported [476- 478]. To resolve this issue, gene reporter assays could be conducted in the presence or absence of inverse agonists that block constitutive receptor activity [479] or downstream inhibitors of β2AR-activated pathways [480], such as PKI (PKA inhibitor), U0126 (MEK kinase inhibitor, prevents ERK1/2 phosphorylation) or dominant negatives p38 mutants (various p38 inhibitors have been shown to directly affect activin signaling activity, [481]). It also remains to be determined if ALK4/5 activation has any direct effect on

β2AR signaling. In this case, variations in the levels of β2AR downstream effectors (cAMP, phosphorylated ERK1/2 or p38) can be measured in cells treated with isoproterenol alone or in the presence of TGFβ ligands (activin/TGFβ). We commonly use LANCE [482] or EPAC [483] assays to detect variations in cAMP, while ERK 1/2 and p38 phosphorylation levels could be evaluated by immunoblot analysis.

194 Conclusions

The results presented provide insights into how activin signaling might be modulated in gonadotropes. ACVR2 is the main type II receptor in LβT2 cells mediating activin A stimulation of Fshb transcription. We showed that ACVR2 is rapidly turned over in gonadotropes, even in the absence of ligand. The mechanism involved in ACVR2 degradation was not fully characterized, but our results suggest that ACVR2-ECD is released from the plasma membrane. In future studies, we will determine whether ACVR2 is indeed shed and the identity of the enzyme mediating this process.

In addition to ACVR2, our results suggest that BMPR2 short isoform (BMPR2-S) functions as an activin receptor in gonadotrope cells to mediate induction of Fshb expression. BMPR2 binds activins with low affinity and this receptor seems to be also rapidly degraded in LβT2 cells. Still, it remains to be determined whether BMPR2-S is normally expressed in gonadotrope cultures and the exact mechanism by which BMPR2- S transduces activin signaling.

When analyzing the oligomerization potential of type I receptors, we found that ALK4 has the potential to heterodimerize with ALK2 and ALK5. Moreover, our BRET results uncovered an unexpected association between ALK4 and ALK5 with β2AR in a heterologous cell system (HEK293 cells). These interactions were confirmed by co- precipitation analysis and preliminary data suggest that β2AR co-expression negatively affect activin and TGFβ signaling even in the absence of adrenergic stimulation. However, additional assays are required to confirm these results.

Although the findings presented here were studied in light of activin responses, they might be relevant for signaling by other TGFβ family members as well. For instance, variations in ACVR2 expression, will also affect the cell response to several BMPs and GDFs, which signal via this receptor. Also, if ACVR2 is shed, the soluble ECD might bind to GDF8 and BMP7, reducing their bioavailability. Moreover, since BMPR2

195 apparently interacts with activins, BMPs and GDFs through the same binding interface, these ligands could cross-inhibit each other while competing for binding to this receptor.

Further experimentation is required to determine if the mechanisms described here operate in other cell lines, as well as, if they play any role in modulation of activin- induced FSH synthesis in vivo. In addition, future investigations will resolve if the type I receptor heterodimers observed in our BRET assays represent functional interactions.

196 Figure legends

Figure 5.1. BMPR2 turns over rapidly in LβT2 cells. A) LβT2 cells were transfected with 1 μg of BMPR2-GFP2 expression vector or pcDNA3.1. After 24 hours, cells were treated for different periods of time with CHX (5 μg/mL) and lysed in RIPA buffer. Expression levels of the type II receptors were evaluated by immunoblotting using anti- GFP2 antibody (BD). (B) Untransfected LβT2 cells were treated for 0 or 6 hours with CHX (5 μg/mL) and lysed in RIPA buffer. Expression of BMPR2 was determined by immunoblotting using anti-BMPR2 (R&D). GAPDH was used as loading control. Shown are the results from a single experiment (n =1) for each panel.

Figure 5.2. Substitution of five residues from the ACVR2 ECD in BMPR2 confers the ability to further enhance activin A signaling. (A) Sequence of human ACVR2B and human BMPR2 protein sequences, showing the five residues from the receptor extracellular domains that were swapped. GenBank numbers are indicated towards the right. (B) Expression of BMPR2 mutants. CHO cells seeded in 6-well plates were transfected with 1 μg of WT or mutant BMPR2-HA expression vectors. Twenty-four hours after transfection, cell surface proteins were biotinylated and whole cell lysates collected for HA immunoprecipitation analysis. Immunoprecipitated proteins were blotted (IB) with HA antibody or using ABC Vectastain reagent to detect biotinylated receptors. GAPDH was used a loading control. The image shown is representative of three independent experiments. (C) Functionality of BMPR2 mutants. LβT2 cells seeded in 48-well plates were transfected with the -1990/+1 mFshb-luc reporter (225 ng/well) along with 50 ng/well of the indicated BMPR2-HA expression constructs. Cells were treated with 25 ng/mL activin A for 24 hours. Data represent mean +/- SEM of 4 independent experiments. Bonferroni multicomparison test, $ p < 0.001.

Figure 5.3. Isoproterenol treatment does not affect activin A-induced SMAD2 phosphorylation in LβT2 cells. HEK293 cells grown in six-well plates, were transfected with the β2AR using Lipofectamine 2000 following manufacturer’s instructions. The next day, cells were placed in serum-free medium and treated for 10 min with isoproterenol (1

197 μM) or vehicle (ascorbic acid), followed by 30 min treatment with activin A (5 ng/ml), TGFβ1 (5 ng/ml) or vehicle (control) as indicated. After ligand treatment, cells were rinsed once with PBS and lysed in RIPA buffer. Total lysates were blotted (IB) for total SMAD2 (Zymed antibody) or with α-phospho-SMAD2 antibody (Cell Signaling). Shown are results from a single experiment (A) and its corresponding quantification (B). Values were normalized against the control condition (cells transfected with pcDNA3.1 and no ligand).

Figure 5.4. Expression of β2AR followed by isoproterenol treatment reduces TGFβ1 induced CAGA-luc reporter activity. HEK293 cells seeded in 48-well plates were transfected with the CAGA-luc reporter and either a β2AR expression vector or pcDNA3.1 using Lipofectamine 2000. The next day, cells were treated with isoproterenol (10 μM) or vehicle (ascorbic acid) along with either 5 ng/ml of TGFβ1 (A) or activin A (B) in serum-free medium, for 24 hours. Data represent the mean of a single experiment. Treatments were performed in triplicate. Values were normalized relative to the control group in which cell were transfected with pcDNA3.1 and treated only with ascorbic acid.

198 Figure 5.1

A

B

199 Figure 5.2

A 72..LVKKGCW---LDDFNCYDRQECV 91 Q13705 AVR2B_HUMAN (AAH96245.1) 79..LVKQGCWSHIGDPQECHYEECVV 101 Q13873 BMPR2_HUMAN (AAH52985.1)

B

C

200 Figure 5.3

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201 Figure 5.4

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