Vitamins & Hormones Volume 85, 2-349 (2011) Activins and Inhibins Edited by: Gerald Litwack ISBN: 978-0-12-385961-7 Series Editors Copyright Contributors Preface Gerald Litwack Chapter One - Activin Receptor-Like Kinase and the Insulin Gene Rie Watanabe Chapter Two - Crystal Structure of Activin Receptor Type IIB Kinase Domain Seungil Han Chapter Three - Activin/Nodal Signaling and Pluripotency Zhenzhi Chng, Ludovic Vallier, Roger Pedersen Chapter Four - Intracrine Signaling Mechanisms of Activin A and TGF-β Olav A. Gressner Chapter Five - Negative Regulation of Activin Signal Transduction Sun-Cheol Choi, Jin-Kwan Han Chapter Six - Antagonism of Activin by Activin Chimeras Uwe Muenster, Radhika Korupolu, Ratindra Rastogi, Jessica Read, Wolfgang H. Fischer Chapter Seven - Activins and Cell Migration Hong-Yo Kang, Chih-Rong Shyr Chapter Eight - The Synthesis and Secretion of Inhibins Kelly L. Walton, Yogeshwar Makanji, David M. Robertson, Craig A. Harrison Chapter Nine - Multifunctional Roles of Activins in the Brain Hiroshi Ageta, Kunihiro Tsuchida Chapter Ten - The Role of Activin/Nodal and Wnt Signaling in Endoderm Formation Catherine Payne, Jason King, David Hay Chapter Eleven - Activin in Glucose Metabolism Osamu Hashimoto, Masayuki Funaba Chapter Twelve - Activin in Humoral Immune Responses Kenji Ogawa, Masayuki Funaba Chapter Thirteen - The Regulation and Functions of Activin and Follistatin in Inflammation and Immunity Mark P. Hedger, Wendy R. Winnall, David J. Phillips, David M. de Kretser Chapter Fourteen - Feedback Regulation by Inhibins A and B of the Pituitary Secretion of Follicle-Stimulating Hormone Yogeshwar Makanji, Craig A. Harrison, David M. Robertson Chapter Fifteen - Activin A in Nonalcoholic Fatty Liver Disease Arne Yndestad, John Willy Haukeland, Tuva B. Dahl, Bente Halvorsen, Pål Aukrust Subject Index
ii iv xiii-xv xvii-xviii 1-27 29-38 39-58 59-77 79-104 105-128 129-148 149-184 185-206
207-216 217-234 235-253
255-297
299-321 323-342 343-349
Former Editors
ROBERT S. HARRIS
KENNETH V. THIMANN
Newton, Massachusetts
University of California Santa Cruz, California
JOHN A. LORRAINE University of Edinburgh Edinburgh, Scotland
PAUL L. MUNSON University of North Carolina Chapel Hill, North Carolina
JOHN GLOVER University of Liverpool Liverpool, England
GERALD D. AURBACH Metabolic Diseases Branch National Institute of Diabetes and Digestive and Kidney Diseases National Institutes of Health Bethesda, Maryland
IRA G. WOOL University of Chicago Chicago, Illinois
EGON DICZFALUSY Karolinska Sjukhuset Stockholm, Sweden
ROBERT OLSEN School of Medicine State University of New York at Stony Brook Stony Brook, New York
DONALD B. MCCORMICK Department of Biochemistry Emory University School of Medicine, Atlanta, Georgia
Cover photo credit: Han, S. Crystal structure of activin receptor type IIB kinase domain. Vitamins and Hormones (2011) 85, pp. 29–38. Academic Press is an imprint of Elsevier 32 Jamestown Road, London, NW1 7BY, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands Linacre House, Jordan Hill, Oxford OX2 8DP, UK 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA First edition 2011 Copyright # 2011 Elsevier Inc. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email:
[email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made ISBN: 978-0-12-385961-7 ISSN: 0083-6729 For information on all Academic Press publications visit our website at elsevierdirect.com Printed and bound in USA 11 12 13 10 9 8 7 6 5 4 3 2 1
CONTRIBUTORS
Hiroshi Ageta Division for Therapies against Intractable Diseases, Institute for Comprehensive Medical Science (ICMS), Fujita Health University, Toyoake, Aichi, Japan Pa˚l Aukrust Research Institute for Internal Medicine, Oslo University Hospital Rikshospitalet, and Faculty of Medicine, University of Oslo; Section of Clinical Immunology and Infectious Diseases, Oslo University Hospital Rikshospitalet, Oslo, Norway Zhenzhi Chng Institute of Medical Biology, Singapore, Singapore Sun-Cheol Choi Division of Molecular and Life Sciences, Pohang University of Science and Technology, Hyoja-dong, Nam-gu, Pohang, Kyungbuk, and Department of Medicine, Graduate School, University of Ulsan, Pungnap-Dong, Songpa-Gu, Seoul, Republic of Korea Tuva B. Dahl Research Institute for Internal Medicine, Oslo University Hospital Rikshospitalet, Oslo, Norway David M. de Kretser Monash Institute of Medical Research, Monash University, Monash Medical Centre, Clayton, and Governor of Victoria, Government House, Melbourne, Victoria, Australia Wolfgang H. Fischer Clayton Foundation Laboratories for Peptide Biology, The Salk Institute for Biological Studies, La Jolla, California, USA Masayuki Funaba Division of Applied Biosciences, Kyoto University Graduate School of Agriculture, Kitashirakawa Oiwakecho, Kyoto, Japan Olav A. Gressner Wisplinghoff Medical Laboratories, Classen-Kappelmann Str. 24, Cologne, Germany Bente Halvorsen Research Institute for Internal Medicine, Oslo University Hospital Rikshospitalet, and Faculty of Medicine, University of Oslo, Oslo, Norway xiii
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Contributors
Jin-Kwan Han Division of Molecular and Life Sciences, Pohang University of Science and Technology, Hyoja-dong, Nam-gu, Pohang, Kyungbuk, Republic of Korea Seungil Han Pfizer Inc., Pfizer Global Research & Development, Groton, Connecticut, USA Craig A. Harrison Prince Henry’s Institute of Medical Research, Clayton, Victoria, Australia Osamu Hashimoto Laboratory of Experimental Animal Science, Faculty of Veterinary Medicine, Kitasato University, School of Veterinary Medicine, Towada, Aomori, Japan John Willy Haukeland Department of Gastroenterology, Oslo University Hospital Aker, Oslo, Norway David Hay MRC Centre for Regenerative Medicine, The University of Edinburgh, Edinburgh, United Kingdom Mark P. Hedger Monash Institute of Medical Research, Monash University, Monash Medical Centre, Clayton, Victoria, Australia Hong-Yo Kang Graduate Institute of Clinical Medical Sciences, and Center for Menopause and Reproductive Research, Chang Gung Memorial Hospital-Kaohsiung Medical Center, Chang Gung University, College of Medicine, Kaohsiung, Taiwan Jason King Roslin Cellab, Roslin Biocentre, Roslin, Midlothian, Scotland, United Kingdom Radhika Korupolu Clayton Foundation Laboratories for Peptide Biology, The Salk Institute for Biological Studies, La Jolla, California, USA Yogeshwar Makanji Prince Henry’s Institute of Medical Research, Clayton, Victoria, Australia Uwe Muenster Clayton Foundation Laboratories for Peptide Biology, The Salk Institute for Biological Studies, La Jolla, California, USA, and Bayer Healthcare, Global Drug Discovery, Pharmaceutical Development, Forschungszentrum Aprath, Wuppertal, Germany Kenji Ogawa Molecular Ligand Discovery Research Team, Chemical Genomics Research Group, ASI, RIKEN, Hirosawa, Wako, Saitama, Japan
Contributors
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Catherine Payne MRC Centre for Regenerative Medicine, The University of Edinburgh, Edinburgh, United Kingdom Roger Pedersen Anne McLaren Laboratory for Regenerative Medicine, University of Cambridge, Cambridge, United Kingdom David J. Phillips Monash Institute of Medical Research, Monash University, Monash Medical Centre, Clayton, and Research Services, La Trobe University, Bundoora, Victoria, Australia Ratindra Rastogi Clayton Foundation Laboratories for Peptide Biology, The Salk Institute for Biological Studies, La Jolla, California, USA Jessica Read Clayton Foundation Laboratories for Peptide Biology, The Salk Institute for Biological Studies, La Jolla, California, USA David M. Robertson Prince Henry’s Institute of Medical Research, Clayton, Victoria, Australia Chih-Rong Shyr Department of Medical Laboratory Science and Biotechnology, China Medical University, and Sex Hormone Research Center, China Medical University Hospital, Taichung, Taiwan Kunihiro Tsuchida Division for Therapies against Intractable Diseases, Institute for Comprehensive Medical Science (ICMS), Fujita Health University, Toyoake, Aichi, Japan Ludovic Vallier Anne McLaren Laboratory for Regenerative Medicine, University of Cambridge, Cambridge, United Kingdom Kelly L. Walton Prince Henry’s Institute of Medical Research, Clayton, Victoria, Australia Rie Watanabe Department of Diabetes and Clinical Nutrition, Kyoto University Graduate School of Medicine, and Laboratory of Infection and Prevention, Department of Biological Responses, Institute for Virus Research, Kyoto University, Kyoto, Japan Wendy R. Winnall Monash Institute of Medical Research, Monash University, Monash Medical Centre, Clayton, Victoria, Australia Arne Yndestad Research Institute for Internal Medicine, Oslo University Hospital Rikshospitalet, Oslo, Norway
PREFACE
Activin and inhibin are dimeric proteins. There are three possible activins, based on the content of bA and bB subunits, and two inhibins, based on the content of a and bA or bB subunits. bA subunit is 32 kDa, and activin A (bA–bA) is about 100 kDa. Activin is synthesized in the gonads, pituitary, and placenta, and its action is to stimulate the synthesis and secretion of follicle-stimulating hormone (FSH) from the anterior pituitary. Inhibin inhibits the synthesis of FSH as well as the secretion of gonadotropicreleasing hormone (GnRH) from the hypothalamus. GnRH acts on the gonadotropic cell of the anterior pituitary to cause the release of FSH. Besides its effect on FSH, activin has activities in cell proliferation, metabolism, differentiation, apoptosis, and others. Inhibin’s effect on FSH is known, but less is known about its functions and the mechanism by which it can inhibit the actions of activin. In this volume, these features of the two hormones are reviewed and the latest information on their characteristics and activities is recorded. Chapter 1, entitled “Activin receptor-like kinase and the insulin gene,” is by R. Watanabe. Chapter 2, an important structural chapter, is entitled “Crystal structure of activin receptor type IIB kinase domain” and authored by S. Han. Chapter 3, entitled “Activin/Nodal signaling and pluripotency,” is authored by Z. Chng, L. Vallier, and R. Pedersen. Chapter 4 is entitled “Intracrine signaling mechanisms of activin A and TGF-b” and is by O. A. Gressner. Chapter 5, “Negative regulation of activin signal transduction,” is offered by S.-C. Choi and J.-K. Han. This is followed by Chapter 6, “Antagonism of activin by activin chimeras,” by U. Muenster, R. Korupolu, R. Rastogi, J. Read, and W. H. Fischer. Chapter 7, “Activins and cell migration,” is authored by H.-Y. Kang and C.-R. Shyr. K. L. Walton, Y. Makanji, D. M. Robertson, and C. A. Harrison contributed Chapter 8, “The synthesis and secretion of inhibins.” Regarding the central nervous system, H. Ageta and K. Tsuchida introduce “Multifunctional roles of activins in the brain.” Following along with the biological functions of activin, C. Payne, J. King, and D. Hay report on “The role activin/nodal and Wnt signaling in endoderm formation.” “Activin in glucose metabolism” is covered by O. Hashimoto and M. Funaba, and K. Ogawa and M. Funaba discuss “Activin in humoral immune responses.” “The regulation and functions of activin and follistatin in inflammation and immunity” is a report by M. P. Hedger, W. R. Winnall, D. J. Phillips, and D. M. de Kretser. Y. Makanji, C. A. Harrison, and D. M. Robertson contribute “Feedback regulation by inhibins A and B of the pituitary secretion of xvii
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follicle-stimulating hormone.” Chapter 15 describes “Activin A in nonalcoholic fatty liver disease” by A. Yndestad, J. W. Haukeland, T. B. Dahl., B. Halvorsen, and P. Aukrust. The figure on the book cover is Fig. 2.1. It shows the type I receptor kinase domain structures. (A) The structure of TbRI in complex with FKBP12. (B) ActRI kinase domain in complex with FKBP12 and dorsomorphin. I appreciate the cooperation of Narmada Thangavelu, Lisa Tickner, and Delsy Retchagar, all of Elsevier, in aspects of the production of this volume. Gerald Litwack October 13, 2010
C H A P T E R
O N E
Activin Receptor-Like Kinase and the Insulin Gene Rie Watanabe1 Contents 2 3 5 5 7 9 9 10 11 12 13 13 14 15 16 16
I. Introduction II. TGF-b Family Receptors: ALK III. Activin Isoforms A, B, and AB A. Activins and ALKs B. Pancreatic endocrine cells IV. Nodal V. Insulin Gene Regulation A. Transcription regulation: A element B. Transcription regulation: GG element C. Transcription regulation: cAMP response element (CRE) D. Transcription regulation: C element E. Transcription regulation: E element F. Transcription regulation: Smad-binding element (SBE) VI. Conclusion Acknowledgments References
Abstract The biological responses of the transforming growth factor-b (TGF-b) superfamily, which includes Activins and Nodal, are induced by activation of a receptor complex and Smads. A type I receptor, which is a component of the complex, is known as an activin receptor-like kinase (ALK); currently seven ALKs (ALK1–ALK7) have been identified in humans. Activins signaling, which is mediated by ALK4 and 7 together with ActRIIA and IIB, plays a critical role in glucose-stimulated insulin secretion, development/neogenesis, and glucose homeostatic control of pancreatic endocrine cells; the insulin gene is regulated by these signaling pathways via ALK7,
Department of Diabetes and Clinical Nutrition, Kyoto University Graduate School of Medicine, Kyoto, Japan Current address: Laboratory of Infection and Prevention, Department of Biological Responses, Institute for Virus Research, Kyoto University, Kyoto, Japan
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Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00001-9
#
2011 Elsevier Inc. All rights reserved.
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which is a receptor for Activins AB and B and Nodal. This review discusses signal transduction of ALKs in pancreatic endocrine cells and the role of ALKs in insulin gene regulation. ß 2011 Elsevier Inc.
I. Introduction The transforming growth factor-b (TGF-b) superfamily, which includes TGF-bs, Activins, Nodal, Inhibins, the bone morphogenetic proteins (BMPs), and growth and differentiation factors (GDFs), regulates a wide variety of cellular processes involving proliferation, differentiation, adhesion, apoptosis, and migration. All TGF-b family members are synthesized as precursor proteins and form dimeric ligands, some of which remain inactive as latent forms by binding to their propeptides, for example, TGF-bs and some GDFs, or as trapped forms by extracellular antagonists, for example, follistatin, which inhibits Activins and noggin and chordin which inhibit some BMPs (Moustakas and Heldin, 2009). On release from these inactive states, the dimeric ligands bind to pairs of membrane receptor serine/threonine kinases, type I (activin receptor-like kinases, ALKs) and type II receptors, promoting the formation of heterotetrameric receptor complexes (Fig. 1.1). Ligand binding induces a link between the constitutively active type II receptors and the dormant type I receptors; when the type II receptor phosphorylates a serine/threonine-rich region, called the GS region, in the cytoplasmic domain of the type I receptor, kinase activity of the type I receptor is stimulated, and ligand-dependent signal transduction then advances. Currently, five type II and seven type I receptors have been identified in mammals. In addition, the TGF-b family ligands also interact with type III receptors: epidermal growth factor–Cripto–FRL1– Cryptic (EGF–CFC)/Cripto, endoglin, and the proteoglycan betaglycan, which are coreceptors and either facilitate or limit the signaling of the receptor kinase. In the absence of the ligand, the small proteins FKBP12 and FKBP12.6 bind to the GS region and maintain the inactive conformation of TGF-b type I receptor by occluding the site of phosphorylation under the TGF-b signaling. The activated type I receptor phosphorylates receptor-regulated Smads (R-Smads) in the cytoplasm; phosphorylated R-Smads associate with common-mediator Smad (Co-Smad), Smad4, and the resulting Smad oligomer is then shuttled into the nucleus. In nucleus, the Smad complexes bind to target genes and regulate their expression together with other transcription factors (Fig. 1.1; Lo¨nn et al., 2009; Massague and Gomis, 2006, Massague et al., 2005; Moustakas and Heldin, 2009; Schmierer and Hill, 2007; Zhang, 2009).
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Latent TGF-b s
Follistatin–Activins
Noggin–BMPs
Activin
BMP
TGF-b
Activin /Nodal
TGF-b
P II
Cripto BMP
PP
P II
I
P Smad2
P II
I
P I P Smad8
P Smad1
P Smad3
P Smad5
Non-Smad pathways
: Smad pathways
Smad4 Smad6 Smad7
Cytoplasm
ad
s
Nucleus P Sm
Smad4 P TFs Smads TFs Element
SBE
Element
Gene expression
Promoter
Figure 1.1 TGF-b, Activin/Nodal, and BMP signaling pathways. Smad pathways are indicated. TGF-b and Activin/Nodal type I receptors phosphorylated Smad2 and 3, and BMP type I receptor phosphorylates Smad1, 5, and 8. The activated R-Smads control gene expressions with various transcription factors (TFs) and Smad4 (Co-Smad) via SBE. Smad6 and Smad7, inhibitory Smads (I-Smads), downregulate the Smad pathways. II, type II receptor; I, type I receptor; - - -, downregulation by I-Smads.
II. TGF-b Family Receptors: ALK In early studies, receptor affinity-labeling analyses using radiolabeled TGF-b revealed TGF-b receptors to comprise three distinct size classes: type I, type II, and type III including proteoglycan betaglycan (Cheifetz et al., 1986; Massague and Like, 1985) and endoglin (Cheifetz et al., 1992); other groups have identified receptors of Activin A (EDF; Hino et al., 1989) and BMP4 (BMP2B; Paralkar et al., 1991) with similar approaches. Expression cloning approaches using degenerate DNA primers (Georgi et al., 1990; Mathews and Vale, 1991) or a probe have identified a number of receptor serine/threonine kinases (Franze´n et al., 1993; Ryde´n et al., 1996; ten Dijke et al., 1993, 1994; Tsuchida et al., 1996), which are type I receptors known as ALKs; seven ALKs, ALK1-7, have been identified in mammals to date.
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Furthermore, many studies identified receptor serine/threonine kinases for TGF-b family members (Attisano et al., 1993; Ebner et al., 1993; He et al., 1993; Kang and Reddi, 1996; Lorentzon et al., 1996; Matsuzaki et al., 1993), type II receptors of TGF-b (Lin et al., 1992), Activins (Attisano et al., 1992; Legerski et al., 1992; Mathews and Vale, 1991; Mathews et al., 1992) and BMPs (Kawabata et al., 1995; Liu et al., 1995; Nohno et al., 1995; Rosenzweig et al., 1995), and type III receptors including endoglin (Gougos and Letarte, 1990; Lo´pez-Casillas et al., 1991; Wang et al., 1991). These findings showed that ALKs include an extracellular ligand-binding region, a single transmembrane domain, intracellular serine/threonine kinase and GS regions, except in type II receptors, in which there is no GS region although they are otherwise structurally similar. ALK1, 2, 3, and 6 are involved in BMP signaling (Miyazono et al., 2010) in combination with the type II receptors BMPR-II (Kawabata et al., 1995; Liu et al., 1995; Nohno et al., 1995; Rosenzweig et al., 1995), ActRIIA (Mathews and Vale, 1991), and IIB (Attisano et al., 1992). ALK1, 2, 3, and 6 activate the R-Smads, Smad1, 5, and 8 (Table 1.1). ALK4 and 7 are stimulated by Activins A, B, and AB, Nodal, and some GDFs together with ActRIIA and IIB (Andersson et al., 2006b, 2008; Reissmann et al., 2001; ten Dijke et al., 1994; Tsuchida et al., 2004); ALK5 is activated by TGF-bs through combination with the type II receptor TbRII (Lin et al., 1992; ten Dijke et al., 1994). GDF8 (Myostatin) and GDF11 bind ActRIIA and IIB together with ALK4 and 5 (Andersson et al., 2006a; Lee et al., 2005; Rebbapragada et al., 2003; Tsuchida et al., 2009). Activated ALK4, 5, and 7 phosphorylate Smad2 and 3. However, increasing evidence shows that TGF-b signaling can also activate Smad1 and 5 in a diversity of cell types in culture (Daly et al., 2008; Finnson et al., 2008; Goumans et al., 2003; Liu et al., 2009), and it has been suggested that a reevaluation of TGF-b family signaling is required by elucidating type I receptor and Smad pathways. In TGF-b signaling through receptor–receptor interactions, TbRII binds with high affinity and is responsible for cooperative recruitment and transphosphorylation of its low-affinity ALK5 pair (Wrana et al., 1992, 1994); ALK5 is predicted to be structurally similar to ALK3 (Harrison et al., 2003) and is anticipated to bind in a mode similar to that of ALK3 (Hart et al., 2002; Lin et al., 2006; Shi and Massague, 2003; Zuniga et al., 2005). Although they are structurally similar, recent analysis of TGF-b and BMP ligands bound to their respective type I and type II receptor ectodomains shows that TGF-b ligands contact both receptors tightly while the evolutionarily more ancient BMPs associate more loosely with their receptors (Groppe et al., 2008). Binding of TGF-b to TbRII creates the interface required for ALK5 recruitment to the complex. Thus, signaling regulation through receptor complexes and downstream molecules is still insufficiently clear.
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Table 1.1 Component regulators in TGF-b family member pathways Type I
Type II
Ligand
R-Smads
I-Smads
Co-Smad
ALK1
BMPR-II/ ActRIIA/ ActRIIB
BMPs
Smad1
Smad6
Smad4
Smad5 Smad8
Smad7
Smad2
Smad7
Smad4
Smad7
Smad4
Smad7
Smad4
Smad6
Smad4
ALK2 ALK3 ALK6 ALK4
ALK7 ALK1 ALK2 ALK3 ALK5 ALK2
ALK3 ALK6 ALK4
ActRIIA/ ActRIIB
Activin A, B, AB/ Nodal
TbR-II
TGF-bs
TbR-II
TGF-bs
BMPR-II/ ActRIIA/ ActRIIB
GDFs
ActRIIA/ ActRIIB
ALK5 ALK7
Smad3 Smad1 Smad5 Smad8 Smad2 Smad3 Smad1
Smad5 Smad8 Smad2
Smad7 Smad7
Smad3
ActRIIA also is known as ActRII. ALK3 and ALK6 denote BMPR-IA and BMPR-IB, respectively. ALK4 is ActR-IB, and ALK5 is TbR-I. Type I, type I receptor; Type II, type II receptor; I-Smads, the inhibitory Smads.
III. Activin Isoforms A, B, and AB A. Activins and ALKs Activins are disulfide-linked homo- or heterodimers of the b subunits of Inhibin/Activin A and B; Activin A (bAbA), Activin B (bBbB), and Activin AB (bAbB) and multifunctional proteins were originally identified as factors in ovarian fluid that stimulated the secretion of follicle stimulating hormone from pituitary cells (Ling et al., 1986; Vale et al., 1986). Activins have potent mesoderm-inducing activity in Xenopus laevis (McDowell and Gurdon, 1999); Nodal is also an authentic mesoderm inducer in many species, including mammals (Shen, 2007). Activins are expressed in a wide variety of tissues, and three isoforms A, B, and AB have been isolated from
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natural sources (Ling et al., 1986; Nakamura et al., 1992; Vale et al., 1986). Activin A and AB have equivalent biological activity levels in various assay systems, whereas the biological activity of native Activin B is significantly lower than those of Activin A and AB (Nakamura et al., 1992). Additional Inhibin/Activin b subunit genes (bC and bE) have been identified in mammals (Fang et al., 1996, 1997; Hashimoto et al., 2002; Ho¨tten et al., 1995; Lau et al., 1996; O’Bryan et al., 2000; Schmitt et al., 1996; Vejda et al., 2002). Gene disruption studies have shown that phenotypes of the Inhibin bAand the Inhibin bB-deficient mouse clearly differ, indicating a lack of functional redundancy between Activins A and B during embryogenesis (Matzuk et al., 1995; Vassalli et al., 1994). Furthermore, replacement of the mature region in the gene of Inhibin bA with the corresponding mature region of Inhibin bB compensates for the Inhibin bA phenotype but evokes additional phenotypes (Brown et al., 2000). These findings indicate that the signalings via Activin A and B have disparate behavior, suggesting a lack of effective compensatory mechanisms. ALK4 and 7 utilize Activins: ALK4 is activated through Activin A, B, and AB, and ALK7 is stimulated by Activin B and AB (Tsuchida et al., 2004). ALK4 is ubiquitously expressed while ALK7 is expressed in embryonic brain (Lorentzon et al., 1996; Tsuchida et al., 1996), adult central nervous system (Kang and Reddi, 1996; Lorentzon et al., 1996; Ryde´n et al., 1996; Tsuchida et al., 1996), prostate (Kang and Reddi, 1996), adipose tissue (Kang and Reddi, 1996; Lorentzon et al., 1996), kidney (Ryde´n et al., 1996; Tsuchida et al., 1996), testis (Tsuchida et al., 1996), gastrointestinal tract (Bondestam et al., 2001; Lorentzon et al., 1996), liver (Lorentzon et al., 1996; Tsuchida et al., 1996), heart (Bondestam et al., 2001), thymus (Lorentzon et al., 1996), coagulating gland (Kang and Reddi, 1996), nasal cavity epithelium (Lorentzon et al., 1996), fetal and adult pancreatic islets (Watanabe et al., 1999), MIN6 (Watanabe et al., 1999), and INS-1 (Zhang et al., 2006) cells. Pancreatic b-cell line MIN6 cells, in which ALK4 and ActRIIB expression is barely detectable and weak, respectively, while ALK7 and ActRIIA are abundantly expressed (Tsuchida et al., 2004; Watanabe et al., 1999), are highly sensitive to Activin AB and modestly to Activin B (Tsuchida et al., 2004). Furthermore, Activin AB and B augment DNA-binding transcriptional activities of Smads in a dose-dependent manner, whereas dominant negative ALK7 (ALK7D/N) expression strongly reduces the activities; glucose-stimulated insulin secretion (GSIS) also is enhanced by Activin AB and B (Tsuchida et al., 2004) but not by Activin A (Shibata et al., 1996; Tsuchida et al., 2004), and it has been shown that Activin AB binds to a combination of ALK7 and ActRIIA in MIN6 cells (Tsuchida et al., 2004). In contrast, in HEK293 and HT22 cells that express ActRIIs and ALK4 but not ALK7 and are highly sensitive to Activin A and AB and
Activin Receptor-Like Kinase and the Insulin Gene
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intermediately to Activin B, it has been shown that ALK7 overexpression induces a dramatic augmentation of Activin B sensitivity for DNA-binding transcriptional activity and that Activin AB modestly enhances the activity, although Activin A sensitivity appears to remain unchanged, indicating that ALK7 is an Activin B-preferring receptor in those cell lines (Tsuchida et al., 2004). In addition, coexpression of ALK7 and ALK7D/N reduces ALK7enhanced activity via Activin AB in HEK293 cells. Taken together, these findings suggest that ALK4 is favored by Activin A; although ALK4 is able to respond to Activin isoforms A as well as B, ALK7 prefers Activin B to Activin A, and Activin AB is more effective than the other two isoforms in ALK4- and ALK7-sensitive signal transduction. However, it remains unknown whether ALK4/Activin B- and ALK4/ Activin AB-mediated signaling mechanisms are similar to that through ALK7/Activins B and AB.
B. Pancreatic endocrine cells Activins A and B are expressed in pancreatic islets (Furukawa et al., 1995; La Rosa et al., 2004; Ogawa et al., 1993, 1995; Tsuchida et al., 2004; Wada et al., 1996; Yasuda et al., 1993) including a-, b-, and d-cells, suggesting an autocrine and/or paracrine system of Activin signals within islets, although their actual secretion levels have not been evaluated. Many studies have shown the importance of Activin signals to physiological functions and development/neogenesis of pancreatic endocrine cells. With regard to glucose homeostasis, it has been found that Activin A stimulates GSIS (Florio et al., 2000; Totsuka et al., 1988; Tsuchida et al., 2004; Verspohl et al., 1993) in a concentration-dependent manner (Florio et al., 2000; Verspohl et al., 1993; Yasuda et al., 1993) mediated by Ca2þ entry (Mogami et al., 1995; Shibata et al., 1993) and counteracted by reduction of extracellular Ca2þ (Shibata et al., 1993). Consistently, MIN6 cells, in which expression of ALK4, the Activin A-preferring receptor, is barely detectable, lack the Activin A effect on GSIS (Shibata et al., 1996; Tsuchida et al., 2004) although Activin AB and B augment GSIS. However, HIT-T15 insulinoma cells, in which ALK7 expression is not detectable (Watanabe et al., 1999), exhibit such Activin A effects (Shibata et al., 1996). These findings indicate that ALK4 plays an essential role in GSIS and Ca2þ-mediated mechanisms via Activin A in pancreatic b-cells, which also suggests a role of a combination of Activin B and ALK7 in the control of GSIS. More recently, it has been shown that Activins have opposite responses to Ca2þ influx in pancreatic islets (Bertolino et al., 2008): Activin A increases glucose-stimulated Ca2þ influx whereas Activin B reduces it. In addition, pancreatic islets show different gene-expression profiles of ALK7, Inhibin bA and Inhibin bB at various glucose concentrations (Bertolino et al., 2008; Zhang et al., 2006), indicating that the extracellular glucose condition regulates the expression of
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the genes of Activins and ALKs related to the glucose homeostasis in pancreatic islets. These findings indicate that in pancreatic islets, Activin A and B exhibit contrary behavior in GSIS control, which may be essential to control glucose homeostasis precisely. Because Activin B in pancreatic b-cells can also stimulate GSIS (Tsuchida et al., 2004), other endocrine cells (e.g., a-cells and d-cells) and/or a novel mechanism may be related to GSIS control of islets, and the Activin AB signal might augment the action of Activins A and B in certain glucose conditions in islets. In addition, recent mutant mice studies have shown that TGF-b family members also play an important role in pancreatic islet functions and glucose homeostasis. It has been found that mice lacking follistatin-like 3 (FSTL3), which is an Activins and GDF8 (Myostatin) antagonist, exhibit an altered metabolic phenotype that includes increased pancreatic islet number and size, improved glucose tolerance and enhanced insulin sensitivity (Mukherjee et al., 2007), while conditional adult overexpression of Smad7, a potent cytoplasmic inhibitor of TGF-bs and Activins signaling, reduces pancreatic insulin content and produces severe hypoinsulinemia (Smart et al., 2006). Mice with attenuated ALK3 (BMPR-IA) signaling in b-cells show decreased expression of key genes involved in insulin gene expression and glucose sensing and develop diabetes due to impaired insulin secretion, and further transgenic expression of BMP4 in b-cells enhances GSIS and glucose clearance (Goulley et al., 2007). On the other hand, it has been demonstrated that TGF-b signaling, which includes Activin A, induces definitive endoderm in mouse and human embryonic stem cells (D’Amour et al., 2005; Kubo et al., 2004; Yasunaga et al., 2005). Activin A and B are able to induce the transformation of embryonic stem cells into insulin-producing cells together with other various stimuli (D’Amour et al., 2006; Jiang et al., 2007; Ku et al., 2004; Ricordi and Edlund, 2008; Shi et al., 2005), and a recent report shows that Activin B is more potent than Activin A in inducing expression of PDX-1, which plays an essential role in the development of pancreas during differentiation of human embryonic stem cells (Frandsen et al., 2007). In addition, in X. laevis, Activin or mature Vg1, a TGF-b-related factor, also induces the expression of XlHbox8, a PDX-1 homolog (Gamer and Wright, 1995; Henry et al., 1996). Early pancreatic-bud explants treated with TGF-b1 in vitro enhance the formation of endocrine cells and inhibit the development of acinar tissue (Sanvito et al., 1994), and further treatment of early buds with follistatin, an Activin antagonist, enhances acinar development while inhibiting that of endocrine cells (Miralles et al., 1998). In dorsal development of the chick pancreas, a notochord signal (comprising Activin B and FGF2) represses sonic hedgehog expression and generates larger insulin-secreting islets (Hebrok et al., 1998), and experiments in Xenopus embryos have shown that transient exposure to Activin and RA can induce pancreas development from isolated animal cap ectoderm (Moriya et al., 2000). Activin A associated with HGF or
Activin Receptor-Like Kinase and the Insulin Gene
9
betacellulin induces the conversion of pancreatic AR42J cells derived from a rat pancreatic acinar carcinoma into insulin-secreting cells (Mashima et al., 1996a,b). Furthermore, developing pancreata of mice lacking the Activin type IIB receptor have severely reduced islet mass but apparently normal acinar tissue (Kim et al., 2000), while transgenic mice with mutated Activin type II receptors have smaller islet area (Shiozaki et al., 1999; Yamaoka et al., 1998), lower survival rate, and lower insulin content in the whole pancreas with impaired glucose tolerance (Yamaoka et al., 1998). Thus, evidence strongly suggests that stimulation by Activins plays a critical role in pancreatic b-cell development and production of insulin-positive cells and b-cell functions. However, the molecular mechanisms by which Activins induce development/neogenesis and regulate b-cell functions remain unclear.
IV. Nodal Nodal signaling also involves ALK4 and 7 together with ActRIIA and IIB. Unlike Activins, however, Nodal signaling requires additional coreceptors from the EGF–CFC protein family such as Cripto to assemble its receptor complexes (Schier and Shen, 2000). Cripto has important roles during development and oncogenesis, and independently binds Nodal and ALK4/7 to promote signaling (Reissmann et al., 2001; Yeo and Whitman, 2001). Recently, it has been shown that Activins signaling is inhibited by Cripto overexpression (Adkins et al., 2003; Gray et al., 2003), and two binding mechanisms have been demonstrated: one involves direct interaction between soluble Cripto and Activin B but not Activin A (Adkins et al., 2003); the other involves type II receptor (IIA and IIB) associated binding between Cripto and Activins A and B (Gray et al., 2003) in blocking Activin signaling. In addition, it has been indicated that Cripto functions as a noncompetitive Activin A antagonist (Kelber et al., 2008). Further experiments are required to elucidate the molecular mechanisms of Cripto function, Nodal signaling and other inhibitory reactions to Activins signaling, and dynamic relations within TGF-b family members and/or cell-to-cell signals could well play an important role in regulation of pancreatic b-cell function and action.
V. Insulin Gene Regulation Insulin is a polypeptide hormone critically involved in the control of glucose homeostasis and is synthesized exclusively in pancreatic islet b-cells by various stimuli. The cloning and sequencing of the human insulin gene
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was reported in 1980 (Bell et al., 1980), and gene mapping studies assigned the human insulin gene to chromosome 11 (p15.5; Harper et al., 1981; Owerbach et al., 1980). Currently, insulin genes have been identified among a number of mammalian species (Steiner et al., 1985; Watanabe et al., 2008). Most animals have a single copy of the insulin gene, whereas in mouse and rat, two nonallelic insulin genes are present (Soares et al., 1985; Steiner et al., 1985). In postnatal life, the insulin gene is expressed almost exclusively in pancreatic b-cells, although low levels of insulin are detected in a number of extrapancreatic tissues (Kojima et al., 2004; Rosenzweig et al., 1980) including brain (Devaskar et al., 1994), thymus ( Jolicoeur et al., 1994; Pugliese et al., 1997; Smith et al., 1997; Vafiadis et al., 1997), lachrymal glands (Cunha et al., 2005), and salivary glands (Vallejo et al., 1984). In thymus that ectopically expresses a broad range of tissue-specific genes for negative selection of autoreactive T cells, variations of insulin expression may be especially relevant to diabetes (Pugliese, 1998). However, there is little understanding of the regulatory sequences and their signaling pathways that control insulin gene expression in non-b-cells, and the role of insulin expression in those cells remains largely unclear. The insulin promoter is located within a region spanning about 400 bp that flanks the transcriptional start site (Edlund et al., 1985; German et al., 1995; Hay and Docherty, 2006; Melloul et al., 2002; Walker et al., 1983). This region contains many ciselements that bind transcription factors, some of which are expressed mainly in pancreatic b-cells and a few other endocrine or neural cell types, while others have widespread tissue distribution (German et al., 1992; Glick et al., 2000; Hay and Docherty, 2006; Qiu et al., 2002; Watanabe et al., 2008). This chapter focuses on representative regulatory elements and a Smad-related element within the human insulin promoter (Fig. 1.2).
A. Transcription regulation: A element A elements containing the core sequence, 50 -TAAT-30 (A1, A3, and A5), bind homeodomain proteins (Rudnick et al., 1994). Among these proteins, the pancreatic and duodenal homeobox factor-1 (PDX-1; Offield et al., 1996), also called IPF1 (Ohlsson et al., 1993), STF-1 (Leonard et al., 1993), IDX-1 (Miller et al., 1994), IUF-1 (Boam and Docherty, 1989), and GSF (Marshak et al., 1996; Melloul et al., 1993) is a well-characterized homeodomain protein expressed in pancreatic islets that plays an essential role in development of pancreas and regulates insulin and somatostatin gene promoters (Liberzon et al., 2004; Ohneda et al., 2000). Recent studies show the glucose-responsive region includes the A3 element (da Silva Xavier et al., 2004; MacFarlane et al., 1994; Marshak et al., 1996; Petersen et al., 1994).
11
Activin Receptor-Like Kinase and the Insulin Gene
(+) PDX1 ATF2
PDX1 -336
ATF2 Smads PDX1 PDX1 MafA BETA2 PDX1 -170
SP1
A5 Core
NRE
C2
-319
G2
E2
A3 CRE1 CRE2
SBE GG2 GG1/A2 C1
-216 Pax4
-58
-87 E1
A1
G1
TATA
-82 c-Jun c-Jun CREM CREM
(-)
Figure 1.2 The major cisacting elements in the human insulin promoter. Transcription factors binding to representative elements are shown. The upper transcription factors upregulate the gene expression (þ), the lower factors downregulate it ().
It was reported that HNF-1a and -1b also bind to the A3 element and stimulate the transactivation in the human insulin promoter (Okita et al., 1999).
B. Transcription regulation: GG element In the human insulin promoter, PDX-1 also responds to the core sequences, 50 -GGAAAT-30 (called the GG elements, GG1 and GG2; Boam et al., 1990; Hay and Docherty, 2006; Le Lay et al., 2004; Tomonari et al., 1999) and regulates expression of the gene. GG1 also has been designated the A2 element (German et al., 1995). GG2 is by far the more conserved, being present in the insulin promoter of all mammals except rodent. The human GG2 element is under positive control of PDX-1 (Le Lay et al., 2004), whereas the corresponding region in the rodent gene is negatively regulated by Nkx2.2, a homeodomain transcription factor of the NK2 class (Cissell et al., 2003), which demonstrates a fundamental difference in the regulation of the human and rodent insulin genes. In the human insulin promoter, although the GG2 element displays a lower PDX-1 binding affinity than A3 and A1 elements in gel mobility shift assays, it is more critical to transcriptional activation in b-cell transfection assays (Le Lay and Stein, 2006). Comparison analyses between the GG elements show that a mutation of the GG1 element drastically decreases the transcriptional activity of the human insulin promoter in MIN6 cells, suggesting that the GG1 element may play the more critical role in b-cell-specific transcriptional activity than the GG2 element (Tomonari et al., 1999). In addition, PDX-1-dependent (Watanabe et al., 2008) and glucose-induced (da Silva Xavier et al., 2004) transactivation of the human insulin promoter is also strongly decreased by a mutation of GG1 element. The early study on transacting factors for GG elements of the human insulin gene by DNase footprint analysis shows transaction of a ubiquitous factor with the GG1 element and of a b-cell-specific factor with the GG2
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element (Boam et al., 1990). It has been shown that the transcription factor binding to the GG1 element interacts with a transcription factor that binds to the adjacent C1 element (Tomonari et al., 1999), which binds the basic leucine zipper (bZIP) factor MafA (Matsuoka et al., 2003), while the GG2 element also contributes to synergistic activation by PDX-1 and MafA (Le Lay and Stein, 2006). Taken together, these findings suggest that both of the GG regulatory elements have a function in insulin expression, and that PDX-1 plays a major role in GG regulation together with proximate transcription factors. However, the signaling mechanisms remain unclear.
C. Transcription regulation: cAMP response element (CRE) In the pancreatic b-cell, glucose (Grill and Cerasi, 1974) and hormones including incretins increase intracellular cAMP (Drucker et al., 1987) and Ca2þ. The human insulin promoter has four CREs, which bind to the CREB/ATF family (Inagaki et al., 1992), and all of these sites are transcriptionally active (Hay et al., 2005; Inagaki et al., 1992). CRE1 and CRE2 are found in the promoter region, CRE3 is in the first exon and CRE4 is in the first intron. Recombinant CREB and ATF2 bind to CRE sites in rat and human insulin promoters (Inagaki et al., 1992; Oetjen et al., 1994), and ChIP analysis demonstrates that CREB binds to mouse insulin 2 promoter (Kuroda et al., 2009), whereas only ATF2 markedly enhances glucose-induced transactivation of the human insulin promoter (Ban et al., 2000). In addition, it also has been shown that siRNA-mediated knockdown of ATF-2 diminishes the stimulatory effects of cAMP-related signaling on insulin promoter activity, suggesting that ATF-2 may be a key regulator of the human insulin promoter (Hay et al., 2007). Furthermore, the c-jun protooncogene product (c-Jun), which was able to form a heterodimer with ATF2 and bind to the CRE site with high affinity (Macgregor et al., 1990), represses cAMP-induced activity of the human insulin promoter (Inagaki et al., 1992). The human insulin promoter also has nine CpG sequences located at positions 357, 345, 234, 206, 180, 135, 102, 69, and 19 bp relative to the transcription start site (Kuroda et al., 2009), and the CpG sites at 206 bp and 180 bp are parts of CRE1 and CRE2, respectively. Methylation of the human insulin promoter also suppresses reporter gene expression, suggesting that DNA methylation/demethylation may play a crucial role in insulin gene regulation by ATF2 and CREB. Indeed, in the mouse Insulin 2 gene, specific methylation of the CpG site in CRE alone suppresses promoter activity, and ChIP analysis shows that methylation increases the binding of methyl-CpG-binding protein 2 (MeCP2) and conversely inhibits the binding of ATF2 and CREB (Kuroda et al., 2009).
Activin Receptor-Like Kinase and the Insulin Gene
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D. Transcription regulation: C element The bZIP protein MafA has been identified as the rat insulin promoter element 3b1 (RIPE3b1) factor (Shieh and Tsai, 1991), which is a transcription factor that binds to the C1/RIPE3b1 element and positively regulates the transcriptional activity of the insulin promoters in mouse (Matsuoka et al., 2003), rat (Kajihara et al., 2003; Matsuoka et al., 2003; Olbrot et al., 2002), and human (Kataoka et al., 2002). The C1/RIPE3b1 element also has been shown to play a critical role in b-cell-specific insulin gene transcription as well as in its glucose-regulated expression (Kataoka et al., 2002; Sharma and Stein, 1994), and the expression and binding of MafA to the C1/RIPE3b1 element is upregulated in a glucose-sensitive manner (Kataoka et al., 2002; Sharma and Stein, 1994; Sharma et al., 1995). It has been found that the transcription factors PDX-1, MafA, and BETA2, which bind to the A3, C1, and E1 elements, respectively, synergistically control glucose-regulated transcription of the insulin gene in rat (Zhao et al., 2005), whereas there is no indication of any synergistic effect between PDX-1, MafA, or BETA2 on the human insulin promoter (Docherty et al., 2005). In addition, the additive effect of PDX-1 and MafA is known (Docherty et al., 2005). In the human insulin gene, ATF2 also enhances glucoseinduced transactivation, and c-Jun, which is able to form a heterodimer with ATF2, represses it (Ban et al., 2000; Inagaki et al., 1992). These finding suggest that cooperative regulation among these transcription factors may play a major role in glucose-dependent transcription of the human insulin gene. Indeed, MafA but not MafB also can heterodimerize with c-Jun (Benkhelifa et al., 1998; Kerppola and Curran, 1994). The human insulin promoter also has the C2 element (Read et al., 1997), the DNA-binding activity of which is regulated in a redox-dependent manner (Cakir and Ballinger, 2005; Sen and Packer, 1996). The C2 element is able to bind PAX4, which negatively regulates transcriptional activity (Campbell et al., 1999). Another member of the Pax gene family, Pax6, the one most closely related to Pax4, has no significant effect on the transcriptional activity of the human insulin gene (Campbell et al., 1999), although Pax6 binds to the C2 element and acts as a transactivator of the rat insulin I promoter (Fujitani et al., 1999; Sander et al., 1997).
E. Transcription regulation: E element In the insulin gene, E elements, sharing the consensus sequence 50 CANNTG-30 (Boam et al., 1990; Crowe and Tsai, 1989; Karlsson et al., 1987, 1989; Ohlsson and Edlund, 1986; Whelan et al., 1989), bind transcription factors (Boam et al., 1990; Moss et al., 1988; Nelson et al., 1990; Ohlsson et al., 1988; Peyton et al., 1994; Walker et al., 1990) of the basic helix-loop-helix (bHLH) class that function as potent transcriptional
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activators of tissue-specific genes by forming heterodimers between both ubiquitous and cell-restricted family members (Dumonteil et al., 1998; Naya et al., 1995). Mutagenesis of the E1 element in the human insulin promoter reduces basal (Docherty et al., 2005) and glucose-induced (da Silva Xavier et al., 2004; Odagiri et al., 1996) transcriptional activity. The heterodimer between ubiquitous E47 and neuroendocrine cell specific BETA2/NeuroD binds to the E1 element and induces transactivation in rat (Dumonteil et al., 1998; Naya et al., 1995). Although the E1 element is highly conserved, in the human insulin promoter the E2 element is the homologous sequence not the consensus sequence (Boam et al., 1990); the human E2 sequence is able to bind the ubiquitous transcription factor USF (Read et al., 1993).
F. Transcription regulation: Smad-binding element (SBE) The structures of Smad2, Smad3, and Smad4 include two conserved domains in the amino (MH1) and carboxyl (MH2) termini, connected by a proline-rich nonconserved linker region (Massague et al., 2005). Smad2 and Smad3 are phosphorylated by ALK4, 5, and 7; the phosphorylated Smad complexes translocate into the nucleus and interact with DNAbinding proteins and coactivators (Fig. 1.1). The MH1 domains of Smad3 and 4 but not that of Smad2 can act directly on the DNA sequence 50 -GTCT-30 or its complement 50 -AGAC-30 , called the SBE. Many Smad-responsive promoter regions contain one or more SBEs, which in many instances contain an extra base as 50 -CAGAC-30 . The human insulin promoter has a highly conserved SBE, 50 -CAGAC0 3 , and Activin AB/B and Nodal signaling pathways, which activate ALK7, induce DNA binding of Smad3 and stimulate the transcriptional activity of the human insulin gene (Watanabe et al., 2008). Mutagenesis of the SBE dramatically reduces ALK7/Smad3-induced transcription (Watanabe et al., 2008), suggesting that the SBE plays a crucial role in human insulin gene expression induced by these signals. In addition, PDX-1 is able to predominantly interact with phosphorylated Smad3, and then bind to the promoter in an Activin AB/Nodal-sensitive manner and synergistically upregulate transactivation; this synergy is completely abolished by mutations of the elements A2/GG1, A3, or SBE (Watanabe et al., 2008; Fig. 1.3). Thus, these findings suggest that association between the cell/tissue-specific transcription factor PDX-1 and ubiquitous factors, at least Smad3, on the insulin promoter specifically controls insulin gene expression via Activin AB/B and Nodal signals.
15
Activin Receptor-Like Kinase and the Insulin Gene
Follistatin–Activins
Activin
Nodal
Activin /Nodal
Pancreatic b -cell II
Smad2
Cripto
P P I Smad3
Smad2
P
Smad2
P
P
P
Smad2
P
Smad3
Smad3
Smad3
P
Smad2
Smad2 Smad4
Cytoplasm
Smad7 PDX-1
Nucleus
PDX-1 Sma P d3 ad2 Sm P PDX-1 Smad3 PDX-1
P
A3
SBE
A2/GG1
Insulin
Promoter
Figure 1.3 Activin AB and Nodal signaling in the pancreatic b-cell. Activated ALK7 phosphorylates Smad2 and 3, and the activated Smads bind to the SBE in the human insulin promoter, resulting in stimulation of transcription of the gene together with PDX-1.
VI. Conclusion Growing evidence demonstrates the importance of TGF-b family member signaling as well as that of Activins in physiological functions and development/neogenesis of pancreatic endocrine cells. The insulin promoter is precisely regulated by various stimuli and complex signalings that control b-cell functions and action. Insulin gene transcription is directly stimulated by combination of Smad2/3 and PDX-1 via Activin AB/Nodalassociated ALK7 signalings in pancreatic b-cells, and the highly conserved SBE within the insulin promoter is related to this process. In human, Activins, Nodal, TGF-bs, and some GDFs utilize Smad2 and 3 in control of many cellular processes, suggesting that this SBE of the insulin gene also may be involved in the various signaling pathways through these family members in pancreatic b-cells.
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ACKNOWLEDGMENTS The authors’ work was supported in part by a Grant-in-Aid for JSPS Fellows and Establishment of International COE for Integration of Transplantation Therapy and Regenerative Medicine from the Ministry of Education, Culture, Sports, Science, and Technology (MEXT), Japan.
REFERENCES Adkins, H. B., Bianco, C., Schiffer, S. G., Rayhorn, P., Zafari, M., Cheung, A. E., Orozco, O., Olson, D., De Luca, A., Chen, L. L., Miatkowski, K., Benjamin, C., et al. (2003). Antibody blockade of the Cripto CFC domain suppresses tumor cell growth in vivo. J. Clin. Invest. 112, 575–587. Andersson, O., Reissmann, E., and Ibanez, C. F. (2006a). Growth differentiation factor 11 signals through the transforming growth factor-beta receptor ALK5 to regionalize the anterior-posterior axis. EMBO Rep. 7, 831–837. Andersson, O., Reissmann, E., Jornvall, H., and Ibanez, C. F. (2006b). Synergistic interaction between Gdf1 and Nodal during anterior axis development. Dev. Biol. 293, 370–381. Andersson, O., Korach-Andre, M., Reissmann, E., Ibanez, C. F., and Bertolino, P. (2008). Growth/differentiation factor 3 signals through ALK7 and regulates accumulation of adipose tissue and diet-induced obesity. Proc. Natl. Acad. Sci. USA 105, 7252–7256. Attisano, L., Wrana, J. L., Cheifetz, S., and Massague, J. (1992). Novel activin receptors: Distinct genes and alternative mRNA splicing generate a repertoire of serine/threonine kinase receptors. Cell 68, 97–108. Attisano, L., Carcamo, J., Ventura, F., Weis, F. M., Massague, J., and Wrana, J. L. (1993). Identification of human activin and TGF beta type I receptors that form heteromeric kinase complexes with type II receptors. Cell 75, 671–680. Ban, N., Yamada, Y., Someya, Y., Ihara, Y., Adachi, T., Kubota, A., Watanabe, R., Kuroe, A., Inada, A., Miyawaki, K., Sunaga, Y., Shen, Z. P., et al. (2000). Activating transcription factor-2 is a positive regulator in CaM kinase IV-induced human insulin gene expression. Diabetes 49, 1142–1148. Bell, G. I., Pictet, R. L., Rutter, W. J., Cordell, B., Tischer, E., and Goodman, H. M. (1980). Sequence of the human insulin gene. Nature 284, 26–32. Benkhelifa, S., Provot, S., Lecoq, O., Pouponnot, C., Calothy, G., and FelderSchmittbuhl, M. P. (1998). mafA, a novel member of the maf proto-oncogene family, displays developmental regulation and mitogenic capacity in avian neuroretina cells. Oncogene 17, 247–254. Bertolino, P., Holmberg, R., Reissmann, E., Andersson, O., Berggren, P. O., and Ibanez, C. F. (2008). Activin B receptor ALK7 is a negative regulator of pancreatic beta-cell function. Proc. Natl. Acad. Sci. USA 105, 7246–7251. Boam, D. S., and Docherty, K. (1989). A tissue-specific nuclear factor binds to multiple sites in the human insulin-gene enhancer. Biochem. J. 264, 233–239. Boam, D. S., Clark, A. R., and Docherty, K. (1990). Positive and negative regulation of the human insulin gene by multiple trans-acting factors. J. Biol. Chem. 265, 8285–8296. Bondestam, J., Huotari, M. A., Moren, A., Ustinov, J., Kaivo-Oja, N., Kallio, J., HorelliKuitunen, N., Aaltonen, J., Fujii, M., Moustakas, A., Ten Dijke, P., Otonkoski, T., et al. (2001). cDNA cloning, expression studies and chromosome mapping of human type I serine/threonine kinase receptor ALK7 (ACVR1C). Cytogenet. Cell Genet. 95, 157–162.
Activin Receptor-Like Kinase and the Insulin Gene
17
Brown, C. W., Houston-Hawkins, D. E., Woodruff, T. K., and Matzuk, M. M. (2000). Insertion of Inhbb into the Inhba locus rescues the Inhba-null phenotype and reveals new activin functions. Nat. Genet. 25, 453–457. Cakir, Y., and Ballinger, S. W. (2005). Reactive species-mediated regulation of cell signaling and the cell cycle: The role of MAPK. Antioxid. Redox Signal. 7, 726–740. Campbell, S. C., Cragg, H., Elrick, L. J., Macfarlane, W. M., Shennan, K. I., and Docherty, K. (1999). Inhibitory effect of pax4 on the human insulin and islet amyloid polypeptide (IAPP) promoters. FEBS Lett. 463, 53–57. Cheifetz, S., Like, B., and Massague, J. (1986). Cellular distribution of type I and type II receptors for transforming growth factor-beta. J. Biol. Chem. 261, 9972–9978. Cheifetz, S., Bellon, T., Cales, C., Vera, S., Bernabeu, C., Massague, J., and Letarte, M. (1992). Endoglin is a component of the transforming growth factor-beta receptor system in human endothelial cells. J. Biol. Chem. 267, 19027–19030. Cissell, M. A., Zhao, L., Sussel, L., Henderson, E., and Stein, R. (2003). Transcription factor occupancy of the insulin gene in vivo. Evidence for direct regulation by Nkx2.2. J. Biol. Chem. 278, 751–756. Crowe, D. T., and Tsai, M. J. (1989). Mutagenesis of the rat insulin II 50 -flanking region defines sequences important for expression in HIT cells. Mol. Cell. Biol. 9, 1784–1789. Cunha, D. A., Carneiro, E. M., Alves Mde, C., Jorge, A. G., de Sousa, S. M., Boschero, A. C., Saad, M. J., Velloso, L. A., and Rocha, E. M. (2005). Insulin secretion by rat lacrimal glands: Effects of systemic and local variables. Am. J. Physiol. Endocrinol. Metab. 289, E768–E775. Daly, A. C., Randall, R. A., and Hill, C. S. (2008). Transforming growth factor betainduced Smad1/5 phosphorylation in epithelial cells is mediated by novel receptor complexes and is essential for anchorage-independent growth. Mol. Cell. Biol. 28, 6889–6902. D’Amour, K. A., Agulnick, A. D., Eliazer, S., Kelly, O. G., Kroon, E., and Baetge, E. E. (2005). Efficient differentiation of human embryonic stem cells to definitive endoderm. Nat. Biotechnol. 23, 1534–1541. D’Amour, K. A., Bang, A. G., Eliazer, S., Kelly, O. G., Agulnick, A. D., Smart, N. G., Moorman, M. A., Kroon, E., Carpenter, M. K., and Baetge, E. E. (2006). Production of pancreatic hormone-expressing endocrine cells from human embryonic stem cells. Nat. Biotechnol. 24, 1392–1401. da Silva Xavier, G., Rutter, J., and Rutter, G. A. (2004). Involvement of Per-Arnt-Sim (PAS) kinase in the stimulation of preproinsulin and pancreatic duodenum homeobox 1 gene expression by glucose. Proc. Natl. Acad. Sci. USA 101, 8319–8324. Devaskar, S. U., Giddings, S. J., Rajakumar, P. A., Carnaghi, L. R., Menon, R. K., and Zahm, D. S. (1994). Insulin gene expression and insulin synthesis in mammalian neuronal cells. J. Biol. Chem. 269, 8445–8454. Docherty, H. M., Hay, C. W., Ferguson, L. A., Barrow, J., Durward, E., and Docherty, K. (2005). Relative contribution of PDX-1, MafA and E47/beta2 to the regulation of the human insulin promoter. Biochem. J. 389, 813–820. Drucker, D. J., Philippe, J., Mojsov, S., Chick, W. L., and Habener, J. F. (1987). Glucagonlike peptide I stimulates insulin gene expression and increases cyclic AMP levels in a rat islet cell line. Proc. Natl. Acad. Sci. USA 84, 3434–3438. Dumonteil, E., Laser, B., Constant, I., and Philippe, J. (1998). Differential regulation of the glucagon and insulin I gene promoters by the basic helix-loop-helix transcription factors E47 and BETA2. J. Biol. Chem. 273, 19945–19954. Ebner, R., Chen, R. H., Shum, L., Lawler, S., Zioncheck, T. F., Lee, A., Lopez, A. R., and Derynck, R. (1993). Cloning of a type I TGF-beta receptor and its effect on TGF-beta binding to the type II receptor. Science 260, 1344–1348.
18
Rie Watanabe
Edlund, T., Walker, M. D., Barr, P. J., and Rutter, W. J. (1985). Cell-specific expression of the rat insulin gene: Evidence for role of two distinct 50 flanking elements. Science 230, 912–916. Fang, J., Yin, W., Smiley, E., Wang, S. Q., and Bonadio, J. (1996). Molecular cloning of the mouse activin beta E subunit gene. Biochem. Biophys. Res. Commun. 228, 669–674. Fang, J., Wang, S. Q., Smiley, E., and Bonadio, J. (1997). Genes coding for mouse activin beta C and beta E are closely linked and exhibit a liver-specific expression pattern in adult tissues. Biochem. Biophys. Res. Commun. 231, 655–661. Finnson, K. W., Parker, W. L., ten Dijke, P., Thorikay, M., and Philip, A. (2008). ALK1 opposes ALK5/Smad3 signaling and expression of extracellular matrix components in human chondrocytes. J. Bone Miner. Res. 23, 896–906. Florio, P., Luisi, S., Marchetti, P., Lupi, R., Cobellis, L., Falaschi, C., Sugino, H., Navalesi, R., Genazzani, A. R., and Petraglia, F. (2000). Activin A stimulates insulin secretion in cultured human pancreatic islets. J. Endocrinol. Invest. 23, 231–234. Frandsen, U., Porneki, A. D., Floridon, C., Abdallah, B. M., and Kassem, M. (2007). Activin B mediated induction of Pdx1 in human embryonic stem cell derived embryoid bodies. Biochem. Biophys. Res. Commun. 362, 568–574. Franze´n, P., ten Dijke, P., Ichijo, H., Yamashita, H., Schulz, P., Heldin, C. H., and Miyazono, K. (1993). Cloning of a TGF beta type I receptor that forms a heteromeric complex with the TGF beta type II receptor. Cell 75, 681–692. Fujitani, Y., Kajimoto, Y., Yasuda, T., Matsuoka, T. A., Kaneto, H., Umayahara, Y., Fujita, N., Watada, H., Miyazaki, J. I., Yamasaki, Y., and Hori, M. (1999). Identification of a portable repression domain and an E1A-responsive activation domain in Pax4: A possible role of Pax4 as a transcriptional repressor in the pancreas. Mol. Cell. Biol. 19, 8281–8291. Furukawa, M., Eto, Y., and Kojima, I. (1995). Expression of immunoreactive activin A in fetal rat pancreas. Endocr. J. 42, 63–68. Gamer, L. W., and Wright, C. V. (1995). Autonomous endodermal determination in Xenopus: Regulation of expression of the pancreatic gene XlHbox 8. Dev. Biol. 171, 240–251. Georgi, L. L., Albert, P. S., and Riddle, D. L. (1990). daf-1, a C. elegans gene controlling dauer larva development, encodes a novel receptor protein kinase. Cell 61, 635–645. German, M. S., Wang, J., Chadwick, R. B., and Rutter, W. J. (1992). Synergistic activation of the insulin gene by a LIM-homeo domain protein and a basic helix-loop-helix protein: Building a functional insulin minienhancer complex. Genes Dev. 6, 2165–2176. German, M., Ashcroft, S., Docherty, K., Edlund, H., Edlund, T., Goodison, S., Imura, H., Kennedy, G., Madsen, O., Melloul, D., et al. (1995). The insulin gene promoter. A simplified nomenclature. Diabetes 44, 1002–1004. Glick, E., Leshkowitz, D., and Walker, M. D. (2000). Transcription factor BETA2 acts cooperatively with E2A and PDX1 to activate the insulin gene promoter. J. Biol. Chem. 275, 2199–2204. Gougos, A., and Letarte, M. (1990). Primary structure of endoglin, an RGD-containing glycoprotein of human endothelial cells. J. Biol. Chem. 265, 8361–8364. Goulley, J., Dahl, U., Baeza, N., Mishina, Y., and Edlund, H. (2007). BMP4-BMPR1A signaling in beta cells is required for and augments glucose-stimulated insulin secretion. Cell Metab. 5, 207–219. Goumans, M. J., Valdimarsdottir, G., Itoh, S., Lebrin, F., Larsson, J., Mummery, C., Karlsson, S., and ten Dijke, P. (2003). Activin receptor-like kinase (ALK)1 is an antagonistic mediator of lateral TGFbeta/ALK5 signaling. Mol. Cell 12, 817–828. Gray, P. C., Harrison, C. A., and Vale, W. (2003). Cripto forms a complex with activin and type II activin receptors and can block activin signaling. Proc. Natl. Acad. Sci. USA 100, 5193–5198.
Activin Receptor-Like Kinase and the Insulin Gene
19
Grill, V., and Cerasi, E. (1974). Stimulation by D-glucose of cyclic adenosine 30 :50 -monophosphate accumulation and insulin release in isolated pancreatic islets of the rat. J. Biol. Chem. 249, 4196–4201. Groppe, J., Hinck, C. S., Samavarchi-Tehrani, P., Zubieta, C., Schuermann, J. P., Taylor, A. B., Schwarz, P. M., Wrana, J. L., and Hinck, A. P. (2008). Cooperative assembly of TGF-beta superfamily signaling complexes is mediated by two disparate mechanisms and distinct modes of receptor binding. Mol. Cell 29, 157–168. Harper, M. E., Ullrich, A., and Saunders, G. F. (1981). Localization of the human insulin gene to the distal end of the short arm of chromosome 11. Proc. Natl. Acad. Sci. USA 78, 4458–4460. Harrison, C. A., Gray, P. C., Koerber, S. C., Fischer, W., and Vale, W. (2003). Identification of a functional binding site for activin on the type I receptor ALK4. J. Biol. Chem. 278, 21129–21135. Hart, P. J., Deep, S., Taylor, A. B., Shu, Z., Hinck, C. S., and Hinck, A. P. (2002). Crystal structure of the human TbetaR2 ectodomain—TGF-beta3 complex. Nat. Struct. Biol. 9, 203–208. Hashimoto, O., Tsuchida, K., Ushiro, Y., Hosoi, Y., Hoshi, N., Sugino, H., and Hasegawa, Y. (2002). cDNA cloning and expression of human activin betaE subunit. Mol. Cell. Endocrinol. 194, 117–122. Hay, C. W., and Docherty, K. (2006). Comparative analysis of insulin gene promoters: Implications for diabetes research. Diabetes 55, 3201–3213. Hay, C. W., Sinclair, E. M., Bermano, G., Durward, E., Tadayyon, M., and Docherty, K. (2005). Glucagon-like peptide-1 stimulates human insulin promoter activity in part through cAMP-responsive elements that lie upstream and downstream of the transcription start site. J. Endocrinol. 186, 353–365. Hay, C. W., Ferguson, L. A., and Docherty, K. (2007). ATF-2 stimulates the human insulin promoter through the conserved CRE2 sequence. Biochim. Biophys. Acta 1769, 79–91. He, W. W., Gustafson, M. L., Hirobe, S., and Donahoe, P. K. (1993). Developmental expression of four novel serine/threonine kinase receptors homologous to the activin/ transforming growth factor-beta type II receptor family. Dev. Dyn. 196, 133–142. Hebrok, M., Kim, S. K., and Melton, D. A. (1998). Notochord repression of endodermal Sonic hedgehog permits pancreas development. Genes Dev. 12, 1705–1713. Henry, G. L., Brivanlou, I. H., Kessler, D. S., Hemmati-Brivanlou, A., and Melton, D. A. (1996). TGF-beta signals and a pattern in Xenopus laevis endodermal development. Development 122, 1007–1015. Hino, M., Tojo, A., Miyazono, K., Miura, Y., Chiba, S., Eto, Y., Shibai, H., and Takaku, F. (1989). Characterization of cellular receptors for erythroid differentiation factor on murine erythroleukemia cells. J. Biol. Chem. 264, 10309–10314. Ho¨tten, G., Neidhardt, H., Schneider, C., and Pohl, J. (1995). Cloning of a new member of the TGF-beta family: A putative new activin beta C chain. Biochem. Biophys. Res. Commun. 206, 608–613. Inagaki, N., Maekawa, T., Sudo, T., Ishii, S., Seino, Y., and Imura, H. (1992). c-Jun represses the human insulin promoter activity that depends on multiple cAMP response elements. Proc. Natl. Acad. Sci. USA 89, 1045–1049. Jiang, W., Shi, Y., Zhao, D., Chen, S., Yong, J., Zhang, J., Qing, T., Sun, X., Zhang, P., Ding, M., Li, D., and Deng, H. (2007). In vitro derivation of functional insulinproducing cells from human embryonic stem cells. Cell Res. 17, 333–344. Jolicoeur, C., Hanahan, D., and Smith, K. M. (1994). T-cell tolerance toward a transgenic beta-cell antigen and transcription of endogenous pancreatic genes in thymus. Proc. Natl. Acad. Sci. USA 91, 6707–6711. Kajihara, M., Sone, H., Amemiya, M., Katoh, Y., Isogai, M., Shimano, H., Yamada, N., and Takahashi, S. (2003). Mouse MafA, homologue of zebrafish somite Maf 1, contributes to
20
Rie Watanabe
the specific transcriptional activity through the insulin promoter. Biochem. Biophys. Res. Commun. 312, 831–842. Kang, Y., and Reddi, A. H. (1996). Identification and cloning of a novel type I serine/ threonine kinase receptor of the TGF-beta/BMP superfamily in rat prostate. Biochem. Mol. Biol. Int. 40, 993–1001. Karlsson, O., Edlund, T., Moss, J. B., Rutter, W. J., and Walker, M. D. (1987). A mutational analysis of the insulin gene transcription control region: Expression in beta cells is dependent on two related sequences within the enhancer. Proc. Natl. Acad. Sci. USA 84, 8819–8823. Karlsson, O., Walker, M. D., Rutter, W. J., and Edlund, T. (1989). Individual proteinbinding domains of the insulin gene enhancer positively activate beta-cell-specific transcription. Mol. Cell. Biol. 9, 823–827. Kataoka, K., Han, S. I., Shioda, S., Hirai, M., Nishizawa, M., and Handa, H. (2002). MafA is a glucose-regulated and pancreatic beta-cell-specific transcriptional activator for the insulin gene. J. Biol. Chem. 277, 49903–49910. Kawabata, M., Chytil, A., and Moses, H. L. (1995). Cloning of a novel type II serine/ threonine kinase receptor through interaction with the type I transforming growth factor-beta receptor. J. Biol. Chem. 270, 5625–5630. Kelber, J. A., Shani, G., Booker, E. C., Vale, W. W., and Gray, P. C. (2008). Cripto is a noncompetitive activin antagonist that forms analogous signaling complexes with activin and nodal. J. Biol. Chem. 283, 4490–4500. Kerppola, T. K., and Curran, T. (1994). A conserved region adjacent to the basic domain is required for recognition of an extended DNA binding site by Maf/Nrl family proteins. Oncogene 9, 3149–3158. Kim, S. K., Hebrok, M., Li, E., Oh, S. P., Schrewe, H., Harmon, E. B., Lee, J. S., and Melton, D. A. (2000). Activin receptor patterning of foregut organogenesis. Genes Dev. 14, 1866–1871. Kojima, H., Fujimiya, M., Matsumura, K., Nakahara, T., Hara, M., and Chan, L. (2004). Extrapancreatic insulin-producing cells in multiple organs in diabetes. Proc. Natl. Acad. Sci. USA 101, 2458–2463. Ku, H. T., Zhang, N., Kubo, A., O’Connor, R., Mao, M., Keller, G., and Bromberg, J. S. (2004). Committing embryonic stem cells to early endocrine pancreas in vitro. Stem Cells 22, 1205–1217. Kubo, A., Shinozaki, K., Shannon, J. M., Kouskoff, V., Kennedy, M., Woo, S., Fehling, H. J., and Keller, G. (2004). Development of definitive endoderm from embryonic stem cells in culture. Development 131, 1651–1662. Kuroda, A., Rauch, T. A., Todorov, I., Ku, H. T., Al-Abdullah, I. H., Kandeel, F., Mullen, Y., Pfeifer, G. P., and Ferreri, K. (2009). Insulin gene expression is regulated by DNA methylation. PLoS ONE 4, e6953. La Rosa, S., Uccella, S., Marchet, S., Capella, C., and Lloyd, R. V. (2004). Localization of Inhibins and Activins in normal endocrine cells and endocrine tumors of the gut and pancreas: An immunohistochemical and in situ hybridization study. J. Histochem. Cytochem. 52, 217–225. Lau, A. L., Nishimori, K., and Matzuk, M. M. (1996). Structural analysis of the mouse activin beta C gene. Biochim. Biophys. Acta 1307, 145–148. Lee, S. J., Reed, L. A., Davies, M. V., Girgenrath, S., Goad, M. E., Tomkinson, K. N., Wright, J. F., Barker, C., Ehrmantraut, G., Holmstrom, J., Trowell, B., Gertz, B., et al. (2005). Regulation of muscle growth by multiple ligands signaling through activin type II receptors. Proc. Natl. Acad. Sci. USA 102, 18117–18122. Legerski, R., Zhou, X., Dresback, J., Eberspaecher, H., McKinney, S., Segarini, P., and de Crombrugghe, B. (1992). Molecular cloning and characterization of a novel rat activin receptor. Biochem. Biophys. Res. Commun. 183, 672–679.
Activin Receptor-Like Kinase and the Insulin Gene
21
Le Lay, J., and Stein, R. (2006). Involvement of PDX-1 in activation of human insulin gene transcription. J. Endocrinol. 188, 287–294. Le Lay, J., Matsuoka, T. A., Henderson, E., and Stein, R. (2004). Identification of a novel PDX-1 binding site in the human insulin gene enhancer. J. Biol. Chem. 279, 22228–22235. Leonard, J., Peers, B., Johnson, T., Ferreri, K., Lee, S., and Montminy, M. R. (1993). Characterization of somatostatin transactivating factor-1, a novel homeobox factor that stimulates somatostatin expression in pancreatic islet cells. Mol. Endocrinol. 7, 1275–1283. Liberzon, A., Ridner, G., and Walker, M. D. (2004). Role of intrinsic DNA binding specificity in defining target genes of the mammalian transcription factor PDX1. Nucleic Acids Res. 32, 54–64. Lin, H. Y., Wang, X. F., Ng-Eaton, E., Weinberg, R. A., and Lodish, H. F. (1992). Expression cloning of the TGF-beta type II receptor, a functional transmembrane serine/threonine kinase. Cell 68, 775–785. Lin, S. J., Lerch, T. F., Cook, R. W., Jardetzky, T. S., and Woodruff, T. K. (2006). The structural basis of TGF-beta, bone morphogenetic protein, and activin ligand binding. Reproduction 132, 179–190. Ling, N., Ying, S. Y., Ueno, N., Shimasaki, S., Esch, F., Hotta, M., and Guillemin, R. (1986). Pituitary FSH is released by a heterodimer of the beta-subunits from the two forms of Inhibin. Nature 321, 779–782. Liu, F., Ventura, F., Doody, J., and Massague, J. (1995). Human type II receptor for bone morphogenic proteins (BMPs): Extension of the two-kinase receptor model to the BMPs. Mol. Cell. Biol. 15, 3479–3486. Liu, I. M., Schilling, S. H., Knouse, K. A., Choy, L., Derynck, R., and Wang, X. F. (2009). TGFbeta-stimulated Smad1/5 phosphorylation requires the ALK5 L45 loop and mediates the pro-migratory TGFbeta switch. EMBO J. 28, 88–98. Lo¨nn, P., More´n, A., Raja, E., Dahl, M., and Moustakas, A. (2009). Regulating the stability of TGFbeta receptors and Smads. Cell Res. 19, 21–35. Lo´pez-Casillas, F., Cheifetz, S., Doody, J., Andres, J. L., Lane, W. S., and Massague, J. (1991). Structure and expression of the membrane proteoglycan betaglycan, a component of the TGF-beta receptor system. Cell 67, 785–795. Lorentzon, M., Hoffer, B., Ebendal, T., Olson, L., and Tomac, A. (1996). Habrec1, a novel serine/threonine kinase TGF-beta type I-like receptor, has a specific cellular expression suggesting function in the developing organism and adult brain. Exp. Neurol. 142, 351–360. MacFarlane, W. M., Read, M. L., Gilligan, M., Bujalska, I., and Docherty, K. (1994). Glucose modulates the binding activity of the beta-cell transcription factor IUF1 in a phosphorylation-dependent manner. Biochem. J. 303(Pt 2), 625–631. Macgregor, P. F., Abate, C., and Curran, T. (1990). Direct cloning of leucine zipper proteins: Jun binds cooperatively to the CRE with CRE-BP1. Oncogene 5, 451–458. Marshak, S., Totary, H., Cerasi, E., and Melloul, D. (1996). Purification of the beta-cell glucose-sensitive factor that transactivates the insulin gene differentially in normal and transformed islet cells. Proc. Natl. Acad. Sci. USA 93, 15057–15062. Mashima, H., Ohnishi, H., Wakabayashi, K., Mine, T., Miyagawa, J., Hanafusa, T., Seno, M., Yamada, H., and Kojima, I. (1996a). Betacellulin and activin A coordinately convert amylase-secreting pancreatic AR42J cells into insulin-secreting cells. J. Clin. Invest. 97, 1647–1654. Mashima, H., Shibata, H., Mine, T., and Kojima, I. (1996b). Formation of insulin-producing cells from pancreatic acinar AR42J cells by hepatocyte growth factor. Endocrinology 137, 3969–3976. Massague, J., and Gomis, R. R. (2006). The logic of TGFbeta signaling. FEBS Lett. 580, 2811–2820.
22
Rie Watanabe
Massague, J., and Like, B. (1985). Cellular receptors for type beta transforming growth factor. Ligand binding and affinity labeling in human and rodent cell lines. J. Biol. Chem. 260, 2636–2645. Massague, J., Seoane, J., and Wotton, D. (2005). Smad transcription factors. Genes Dev. 19, 2783–2810. Mathews, L. S., and Vale, W. W. (1991). Expression cloning of an activin receptor, a predicted transmembrane serine kinase. Cell 65, 973–982. Mathews, L. S., Vale, W. W., and Kintner, C. R. (1992). Cloning of a second type of activin receptor and functional characterization in Xenopus embryos. Science 255, 1702–1705. Matsuoka, T. A., Zhao, L., Artner, I., Jarrett, H. W., Friedman, D., Means, A., and Stein, R. (2003). Members of the large Maf transcription family regulate insulin gene transcription in islet beta cells. Mol. Cell. Biol. 23, 6049–6062. Matsuzaki, K., Xu, J., Wang, F., McKeehan, W. L., Krummen, L., and Kan, M. (1993). A widely expressed transmembrane serine/threonine kinase that does not bind activin, inhibin, transforming growth factor beta, or bone morphogenic factor. J. Biol. Chem. 268, 12719–12723. Matzuk, M. M., Kumar, T. R., Vassalli, A., Bickenbach, J. R., Roop, D. R., Jaenisch, R., and Bradley, A. (1995). Functional analysis of activins during mammalian development. Nature 374, 354–356. McDowell, N., and Gurdon, J. B. (1999). Activin as a morphogen in Xenopus mesoderm induction. Semin. Cell Dev. Biol. 10, 311–317. Melloul, D., Ben-Neriah, Y., and Cerasi, E. (1993). Glucose modulates the binding of an islet-specific factor to a conserved sequence within the rat I and the human insulin promoters. Proc. Natl. Acad. Sci. USA 90, 3865–3869. Melloul, D., Marshak, S., and Cerasi, E. (2002). Regulation of insulin gene transcription. Diabetologia 45, 309–326. Miller, C. P., McGehee, R. E., Jr., and Habener, J. F. (1994). IDX-1: A new homeodomain transcription factor expressed in rat pancreatic islets and duodenum that transactivates the somatostatin gene. EMBO J. 13, 1145–1156. Miralles, F., Czernichow, P., and Scharfmann, R. (1998). Follistatin regulates the relative proportions of endocrine versus exocrine tissue during pancreatic development. Development 125, 1017–1024. Miyazono, K., Kamiya, Y., and Morikawa, M. (2010). Bone morphogenetic protein receptors and signal transduction. J. Biochem. 147, 35–51. Mogami, H., Kanzaki, M., Nobusawa, R., Zhang, Y. Q., Furukawa, M., and Kojima, I. (1995). Modulation of adenosine triphosphate-sensitive potassium channel and voltage-dependent calcium channel by activin A in HIT-T15 cells. Endocrinology 136, 2960–2966. Moriya, N., Komazaki, S., Takahashi, S., Yokota, C., and Asashima, M. (2000). In vitro pancreas formation from Xenopus ectoderm treated with activin and retinoic acid. Dev. Growth Differ. 42, 593–602. Moss, L. G., Moss, J. B., and Rutter, W. J. (1988). Systematic binding analysis of the insulin gene transcription control region: Insulin and immunoglobulin enhancers utilize similar transactivators. Mol. Cell. Biol. 8, 2620–2627. Moustakas, A., and Heldin, C. H. (2009). The regulation of TGFbeta signal transduction. Development 136, 3699–3714. Mukherjee, A., Sidis, Y., Mahan, A., Raher, M. J., Xia, Y., Rosen, E. D., Bloch, K. D., Thomas, M. K., and Schneyer, A. L. (2007). FSTL3 deletion reveals roles for TGF-beta family ligands in glucose and fat homeostasis in adults. Proc. Natl. Acad. Sci. USA 104, 1348–1353. Nakamura, T., Asashima, M., Eto, Y., Takio, K., Uchiyama, H., Moriya, N., Ariizumi, T., Yashiro, T., Sugino, K., Titani, K., et al. (1992). Isolation and characterization of native activin B. J. Biol. Chem. 267, 16385–16389.
Activin Receptor-Like Kinase and the Insulin Gene
23
Naya, F. J., Stellrecht, C. M., and Tsai, M. J. (1995). Tissue-specific regulation of the insulin gene by a novel basic helix-loop-helix transcription factor. Genes Dev. 9, 1009–1019. Nelson, C., Shen, L. P., Meister, A., Fodor, E., and Rutter, W. J. (1990). Pan: A transcriptional regulator that binds chymotrypsin, insulin, and AP-4 enhancer motifs. Genes Dev. 4, 1035–1043. Nohno, T., Ishikawa, T., Saito, T., Hosokawa, K., Noji, S., Wolsing, D. H., and Rosenbaum, J. S. (1995). Identification of a human type II receptor for bone morphogenetic protein-4 that forms differential heteromeric complexes with bone morphogenetic protein type I receptors. J. Biol. Chem. 270, 22522–22526. O’Bryan, M. K., Sebire, K. L., Gerdprasert, O., Hedger, M. P., Hearn, M. T., and de Kretser, D. M. (2000). Cloning and regulation of the rat activin betaE subunit. J. Mol. Endocrinol. 24, 409–418. Odagiri, H., Wang, J., and German, M. S. (1996). Function of the human insulin promoter in primary cultured islet cells. J. Biol. Chem. 271, 1909–1915. Oetjen, E., Diedrich, T., Eggers, A., Eckert, B., and Knepel, W. (1994). Distinct properties of the cAMP-responsive element of the rat insulin I gene. J. Biol. Chem. 269, 27036–27044. Offield, M. F., Jetton, T. L., Labosky, P. A., Ray, M., Stein, R. W., Magnuson, M. A., Hogan, B. L., and Wright, C. V. (1996). PDX-1 is required for pancreatic outgrowth and differentiation of the rostral duodenum. Development 122, 983–995. Ogawa, K., Abe, K., Kurosawa, N., Kurohmaru, M., Sugino, H., Takahashi, M., and Hayashi, Y. (1993). Expression of alpha, beta A and beta B subunits of inhibin or activin and follistatin in rat pancreatic islets. FEBS Lett. 319, 217–220. Ogawa, K., Ono, K., Kurohmaru, M., and Hayashi, Y. (1995). Effect of streptozotocin injection on expression of immunoreactive follistatin and beta A and beta B subunits of inhibin/activin in rat pancreatic islets. Eur. J. Endocrinol. 132, 363–369. Ohlsson, H., and Edlund, T. (1986). Sequence-specific interactions of nuclear factors with the insulin gene enhancer. Cell 45, 35–44. Ohlsson, H., Karlsson, O., and Edlund, T. (1988). A beta-cell-specific protein binds to the two major regulatory sequences of the insulin gene enhancer. Proc. Natl. Acad. Sci. USA 85, 4228–4231. Ohlsson, H., Karlsson, K., and Edlund, T. (1993). IPF1, a homeodomain-containing transactivator of the insulin gene. EMBO J. 12, 4251–4259. Ohneda, K., Mirmira, R. G., Wang, J., Johnson, J. D., and German, M. S. (2000). The homeodomain of PDX-1 mediates multiple protein-protein interactions in the formation of a transcriptional activation complex on the insulin promoter. Mol. Cell Biol. 20, 900–911. Okita, K., Yang, Q., Yamagata, K., Hangenfeldt, K. A., Miyagawa, J., Kajimoto, Y., Nakajima, H., Namba, M., Wollheim, C. B., Hanafusa, T., and Matsuzawa, Y. (1999). Human insulin gene is a target gene of hepatocyte nuclear factor-1alpha (HNF-1alpha) and HNF-1beta. Biochem. Biophys. Res. Commun. 263, 566–569. Olbrot, M., Rud, J., Moss, L. G., and Sharma, A. (2002). Identification of beta-cell-specific insulin gene transcription factor RIPE3b1 as mammalian MafA. Proc. Natl. Acad. Sci. USA 99, 6737–6742. Owerbach, D., Bell, G. I., Rutter, W. J., and Shows, T. B. (1980). The insulin gene is located on chromosome 11 in humans. Nature 286, 82–84. Paralkar, V. M., Hammonds, R. G., and Reddi, A. H. (1991). Identification and characterization of cellular binding proteins (receptors) for recombinant human bone morphogenetic protein 2B, an initiator of bone differentiation cascade. Proc. Natl. Acad. Sci. USA 88, 3397–3401. Petersen, H. V., Serup, P., Leonard, J., Michelsen, B. K., and Madsen, O. D. (1994). Transcriptional regulation of the human insulin gene is dependent on the homeodomain
24
Rie Watanabe
protein STF1/IPF1 acting through the CT boxes. Proc. Natl. Acad. Sci. USA 91, 10465–10469. Peyton, M., Moss, L. G., and Tsai, M. J. (1994). Two distinct class A helix-loop-helix transcription factors, E2A and BETA1, form separate DNA binding complexes on the insulin gene E box. J. Biol. Chem. 269, 25936–25941. Pugliese, A. (1998). Insulin expression in the thymus, tolerance, and type 1 diabetes. Diabetes Metab. Rev. 14, 325–327. Pugliese, A., Zeller, M., Fernandez, A., Jr., Zalcberg, L. J., Bartlett, R. J., Ricordi, C., Pietropaolo, M., Eisenbarth, G. S., Bennett, S. T., and Patel, D. D. (1997). The insulin gene is transcribed in the human thymus and transcription levels correlated with allelic variation at the INS VNTR-IDDM2 susceptibility locus for type 1 diabetes. Nat. Genet. 15, 293–297. Qiu, Y., Guo, M., Huang, S., and Stein, R. (2002). Insulin gene transcription is mediated by interactions between the p300 coactivator and PDX-1, BETA2, and E47. Mol. Cell. Biol. 22, 412–420. Read, M. L., Clark, A. R., and Docherty, K. (1993). The helix-loop-helix transcription factor USF (upstream stimulating factor) binds to a regulatory sequence of the human insulin gene enhancer. Biochem. J. 295(Pt 1), 233–237. Read, M. L., Masson, M. R., and Docherty, K. (1997). A RIPE3b1-like factor binds to a novel site in the human insulin promoter in a redox-dependent manner. FEBS Lett. 418, 68–72. Rebbapragada, A., Benchabane, H., Wrana, J. L., Celeste, A. J., and Attisano, L. (2003). Myostatin signals through a transforming growth factor beta-like signaling pathway to block adipogenesis. Mol. Cell. Biol. 23, 7230–7242. Reissmann, E., Jornvall, H., Blokzijl, A., Andersson, O., Chang, C., Minchiotti, G., Persico, M. G., Ibanez, C. F., and Brivanlou, A. H. (2001). The orphan receptor ALK7 and the Activin receptor ALK4 mediate signaling by Nodal proteins during vertebrate development. Genes Dev. 15, 2010–2022. Ricordi, C., and Edlund, H. (2008). Toward a renewable source of pancreatic beta-cells. Nat. Biotechnol. 26, 397–398. Rosenzweig, J. L., Havrankova, J., Lesniak, M. A., Brownstein, M., and Roth, J. (1980). Insulin is ubiquitous in extrapancreatic tissues of rats and humans. Proc. Natl. Acad. Sci. USA 77, 572–576. Rosenzweig, B. L., Imamura, T., Okadome, T., Cox, G. N., Yamashita, H., ten Dijke, P., Heldin, C. H., and Miyazono, K. (1995). Cloning and characterization of a human type II receptor for bone morphogenetic proteins. Proc. Natl. Acad. Sci. USA 92, 7632–7636. Rudnick, A., Ling, T. Y., Odagiri, H., Rutter, W. J., and German, M. S. (1994). Pancreatic beta cells express a diverse set of homeobox genes. Proc. Natl. Acad. Sci. USA 91, 12203–12207. Ryde´n, M., Imamura, T., Jornvall, H., Belluardo, N., Neveu, I., Trupp, M., Okadome, T., ten Dijke, P., and Ibanez, C. F. (1996). A novel type I receptor serine-threonine kinase predominantly expressed in the adult central nervous system. J. Biol. Chem. 271, 30603–30609. Sander, M., Neubuser, A., Kalamaras, J., Ee, H. C., Martin, G. R., and German, M. S. (1997). Genetic analysis reveals that PAX6 is required for normal transcription of pancreatic hormone genes and islet development. Genes Dev. 11, 1662–1673. Sanvito, F., Herrera, P. L., Huarte, J., Nichols, A., Montesano, R., Orci, L., and Vassalli, J. D. (1994). TGF-beta 1 influences the relative development of the exocrine and endocrine pancreas in vitro. Development 120, 3451–3462. Schier, A. F., and Shen, M. M. (2000). Nodal signalling in vertebrate development. Nature 403, 385–389. Schmierer, B., and Hill, C. S. (2007). TGFbeta-SMAD signal transduction: Molecular specificity and functional flexibility. Nat. Rev. Mol. Cell Biol. 8, 970–982.
Activin Receptor-Like Kinase and the Insulin Gene
25
Schmitt, J., Hotten, G., Jenkins, N. A., Gilbert, D. J., Copeland, N. G., Pohl, J., and Schrewe, H. (1996). Structure, chromosomal localization, and expression analysis of the mouse inhibin/activin beta C (Inhbc) gene. Genomics 32, 358–366. Sen, C. K., and Packer, L. (1996). Antioxidant and redox regulation of gene transcription. FASEB J. 10, 709–720. Sharma, A., and Stein, R. (1994). Glucose-induced transcription of the insulin gene is mediated by factors required for beta-cell-type-specific expression. Mol. Cell. Biol. 14, 871–879. Sharma, A., Fusco-DeMane, D., Henderson, E., Efrat, S., and Stein, R. (1995). The role of the insulin control element and RIPE3b1 activators in glucose-stimulated transcription of the insulin gene. Mol. Endocrinol. 9, 1468–1476. Shen, M. M. (2007). Nodal signaling: developmental roles and regulation. Development 134, 1023–1034. Shi, Y., and Massague, J. (2003). Mechanisms of TGF-beta signaling from cell membrane to the nucleus. Cell 113, 685–700. Shi, Y., Hou, L., Tang, F., Jiang, W., Wang, P., Ding, M., and Deng, H. (2005). Inducing embryonic stem cells to differentiate into pancreatic beta cells by a novel three-step approach with activin A and all-trans retinoic acid. Stem Cells 23, 656–662. Shibata, H., Yasuda, H., Sekine, N., Mine, T., Totsuka, Y., and Kojima, I. (1993). Activin A increases intracellular free calcium concentrations in rat pancreatic islets. FEBS Lett. 329, 194–198. Shibata, H., Kanzaki, M., Takeuchi, T., Miyazaki, J., and Kojima, I. (1996). Two distinct signaling pathways activated by activin A in glucose-responsive pancreatic beta-cell lines. J. Mol. Endocrinol. 16, 249–258. Shieh, S. Y., and Tsai, M. J. (1991). Cell-specific and ubiquitous factors are responsible for the enhancer activity of the rat insulin II gene. J. Biol. Chem. 266, 16708–16714. Shiozaki, S., Tajima, T., Zhang, Y. Q., Furukawa, M., Nakazato, Y., and Kojima, I. (1999). Impaired differentiation of endocrine and exocrine cells of the pancreas in transgenic mouse expressing the truncated type II activin receptor. Biochim. Biophys. Acta 1450, 1–11. Smart, N. G., Apelqvist, A. A., Gu, X., Harmon, E. B., Topper, J. N., MacDonald, R. J., and Kim, S. K. (2006). Conditional expression of Smad7 in pancreatic beta cells disrupts TGF-beta signaling and induces reversible diabetes mellitus. PLoS Biol. 4, e39. Smith, K. M., Olson, D. C., Hirose, R., and Hanahan, D. (1997). Pancreatic gene expression in rare cells of thymic medulla: Evidence for functional contribution to T cell tolerance. Int. Immunol. 9, 1355–1365. Soares, M. B., Schon, E., Henderson, A., Karathanasis, S. K., Cate, R., Zeitlin, S., Chirgwin, J., and Efstratiadis, A. (1985). RNA-mediated gene duplication: The rat preproinsulin I gene is a functional retroposon. Mol. Cell. Biol. 5, 2090–2103. Steiner, D. F., Chan, S. J., Welsh, J. M., and Kwok, S. C. (1985). Structure and evolution of the insulin gene. Annu. Rev. Genet. 19, 463–484. ten Dijke, P., Ichijo, H., Franze´n, P., Schulz, P., Saras, J., Toyoshima, H., Heldin, C. H., and Miyazono, K. (1993). Activin receptor-like kinases: A novel subclass of cell-surface receptors with predicted serine/threonine kinase activity. Oncogene 8, 2879–2887. ten Dijke, P., Yamashita, H., Ichijo, H., Franze´n, P., Laiho, M., Miyazono, K., and Heldin, C. H. (1994). Characterization of type I receptors for transforming growth factor-beta and activin. Science 264, 101–104. Tomonari, A., Yoshimoto, K., Mizusawa, N., Iwahana, H., and Itakura, M. (1999). Differential regulation of the human insulin gene transcription by GG1 and GG2 elements with GG- and C1-binding factors. Biochim. Biophys. Acta 1446, 233–242. Totsuka, Y., Tabuchi, M., Kojima, I., Shibai, H., and Ogata, E. (1988). A novel action of activin A: Stimulation of insulin secretion in rat pancreatic islets. Biochem. Biophys. Res. Commun. 156, 335–339.
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Tsuchida, K., Sawchenko, P. E., Nishikawa, S., and Vale, W. W. (1996). Molecular cloning of a novel type I receptor serine/threonine kinase for the TGF beta superfamily from rat brain. Mol. Cell. Neurosci. 7, 467–478. Tsuchida, K., Nakatani, M., Yamakawa, N., Hashimoto, O., Hasegawa, Y., and Sugino, H. (2004). Activin isoforms signal through type I receptor serine/threonine kinase ALK7. Mol. Cell. Endocrinol. 220, 59–65. Tsuchida, K., Nakatani, M., Hitachi, K., Uezumi, A., Sunada, Y., Ageta, H., and Inokuchi, K. (2009). Activin signaling as an emerging target for therapeutic interventions. Cell Commun Signal 7, 15. Vafiadis, P., Bennett, S. T., Todd, J. A., Nadeau, J., Grabs, R., Goodyer, C. G., Wickramasinghe, S., Colle, E., and Polychronakos, C. (1997). Insulin expression in human thymus is modulated by INS VNTR alleles at the IDDM2 locus. Nat. Genet. 15, 289–292. Vale, W., Rivier, J., Vaughan, J., McClintock, R., Corrigan, A., Woo, W., Karr, D., and Spiess, J. (1986). Purification and characterization of an FSH releasing protein from porcine ovarian follicular fluid. Nature 321, 776–779. Vallejo, G., Mead, P. M., Gaynor, D. H., Devlin, J. T., and Robbins, D. C. (1984). Characterization of immunoreactive insulin in human saliva: Evidence against production in situ. Diabetologia 27, 437–440. Vassalli, A., Matzuk, M. M., Gardner, H. A., Lee, K. F., and Jaenisch, R. (1994). Activin/ inhibin beta B subunit gene disruption leads to defects in eyelid development and female reproduction. Genes Dev. 8, 414–427. Vejda, S., Cranfield, M., Peter, B., Mellor, S. L., Groome, N., Schulte-Hermann, R., and Rossmanith, W. (2002). Expression and dimerization of the rat activin subunits betaC and betaE: Evidence for the ormation of novel activin dimers. J. Mol. Endocrinol. 28, 137–148. Verspohl, E. J., Ammon, H. P., and Wahl, M. A. (1993). Activin A: Its effects on rat pancreatic islets and the mechanism of action involved. Life Sci. 53, 1069–1078. Wada, M., Shintani, Y., Kosaka, M., Sano, T., Hizawa, K., and Saito, S. (1996). Immunohistochemical localization of activin A and follistatin in human tissues. Endocr. J. 43, 375–385. Walker, M. D., Edlund, T., Boulet, A. M., and Rutter, W. J. (1983). Cell-specific expression controlled by the 50 -flanking region of insulin and chymotrypsin genes. Nature 306, 557–561. Walker, M. D., Park, C. W., Rosen, A., and Aronheim, A. (1990). A cDNA from a mouse pancreatic beta cell encoding a putative transcription factor of the insulin gene. Nucleic Acids Res. 18, 1159–1166. Wang, X. F., Lin, H. Y., Ng-Eaton, E., Downward, J., Lodish, H. F., and Weinberg, R. A. (1991). Expression cloning and characterization of the TGF-beta type III receptor. Cell 67, 797–805. Watanabe, R., Yamada, Y., Ihara, Y., Someya, Y., Kubota, A., Kagimoto, S., Kuroe, A., Iwakura, T., Shen, Z. P., Inada, A., Adachi, T., Ban, N., et al. (1999). The MH1 domains of smad2 and smad3 are involved in the regulation of the ALK7 signals. Biochem. Biophys. Res. Commun. 254, 707–712. Watanabe, R., Shen, Z. P., Tsuda, K., and Yamada, Y. (2008). Insulin gene is a target in activin receptor-like kinase 7 signaling pathway in pancreatic beta-cells. Biochem. Biophys. Res. Commun. 377, 867–872. Whelan, J., Poon, D., Weil, P. A., and Stein, R. (1989). Pancreatic beta-cell-type-specific expression of the rat insulin II gene is controlled by positive and negative cellular transcriptional elements. Mol. Cell. Biol. 9, 3253–3259.
Activin Receptor-Like Kinase and the Insulin Gene
27
Wrana, J. L., Attisano, L., Carcamo, J., Zentella, A., Doody, J., Laiho, M., Wang, X. F., and Massague, J. (1992). TGF beta signals through a heteromeric protein kinase receptor complex. Cell 71, 1003–1014. Wrana, J. L., Attisano, L., Wieser, R., Ventura, F., and Massague, J. (1994). Mechanism of activation of the TGF-beta receptor. Nature 370, 341–347. Yamaoka, T., Idehara, C., Yano, M., Matsushita, T., Yamada, T., Ii, S., Moritani, M., Hata, J., Sugino, H., Noji, S., and Itakura, M. (1998). Hypoplasia of pancreatic islets in transgenic mice expressing activin receptor mutants. J. Clin. Invest. 102, 294–301. Yasuda, H., Inoue, K., Shibata, H., Takeuchi, T., Eto, Y., Hasegawa, Y., Sekine, N., Totsuka, Y., Mine, T., Ogata, E., et al. (1993). Existence of activin-A in A- and D-cells of rat pancreatic islet. Endocrinology 133, 624–630. Yasunaga, M., Tada, S., Torikai-Nishikawa, S., Nakano, Y., Okada, M., Jakt, L. M., Nishikawa, S., Chiba, T., and Era, T. (2005). Induction and monitoring of definitive and visceral endoderm differentiation of mouse ES cells. Nat. Biotechnol. 23, 1542–1550. Yeo, C., and Whitman, M. (2001). Nodal signals to Smads through Cripto-dependent and Cripto-independent mechanisms. Mol. Cell 7, 949–957. Zhang, Y. E. (2009). Non-Smad pathways in TGF-beta signaling. Cell Res. 19, 128–139. Zhang, N., Kumar, M., Xu, G., Ju, W., Yoon, T., Xu, E., Huang, X., Gaisano, H., Peng, C., and Wang, Q. (2006). Activin receptor-like kinase 7 induces apoptosis of pancreatic beta cells and beta cell lines. Diabetologia 49, 506–518. Zhao, L., Guo, M., Matsuoka, T. A., Hagman, D. K., Parazzoli, S. D., Poitout, V., and Stein, R. (2005). The islet beta cell-enriched MafA activator is a key regulator of insulin gene transcription. J. Biol. Chem. 280, 11887–11894. Zuniga, J. E., Groppe, J. C., Cui, Y., Hinck, C. S., Contreras-Shannon, V., Pakhomova, O. N., Yang, J., Tang, Y., Mendoza, V., Lo´pez-Casillas, F., Sun, L., and Hinck, A. P. (2005). Assembly of TbetaRI:TbetaRII:TGFbeta ternary complex in vitro with receptor extracellular domains is cooperative and isoform-dependent. J. Mol. Biol. 354, 1052–1068.
C H A P T E R
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Crystal Structure of Activin Receptor Type IIB Kinase Domain Seungil Han Contents I. Introduction II. Type I Receptor Kinase Domain Structures A. TGF-b RI kinase domain structure B. ActRIA structure in complex with FKPB12 and dorsomorphin III. 3D Structure of ActRIIB A. Overall structure B. Active site C. Comparison to type I/II receptor kinase domain structures IV. Conclusion References
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Abstract Activin receptor type IIB (ActRIIB) belongs to a type II transforming growth factor-b (TGF-b) serine/threonine kinase receptor family which is integral to the activin and myostatin signaling pathway. Actvin and myostatin bind to activin type II receptors (ActRIIA and ActRIIB), and the glycine–serine-rich domains of type I receptors are phosphorylated by type II receptors. Activin enhances follicle-stimulating hormone biosynthesis and secretion and is involved in apoptosis, fibrosis, inflammation, and neurogenesis. Because of its essential role, activin is regarded as a novel drug target. Myostatin, also referred as growth and differentiation factor 8 (GDF-8), modulates skeletal muscle growth and has been a therapeutic target for disease conditions such as muscular dystrophy, sarcopenia, cashexia, and diabetes mellitus. The AcRIIB kinase domain from human represents a distinct type II receptor serine/threonine kinase subfamily identifiable in part by common features of Thr265 as a gatekeeper residue and back pocket supported by Phe247. The human ActRII kinase domain structure provides a basis for a more integrated understanding of substrate recognition and catalysis and will also be of help for developing chemical inhibitors. ß 2011 Elsevier Inc. Pfizer Inc., Pfizer Global Research & Development, Groton, Connecticut, USA Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00002-0
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2011 Elsevier Inc. All rights reserved.
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I. Introduction Activin, inhibin, and myostatin are members of the transforming growth factor-b (TGF-b) family, the members of which have a wide range of biological actions on cell growth and differentiation (McPherron et al., 1997; Sugino and Tsuchida, 2000). Inhibin inhibits secretion of folliclestimulating hormones (FSHs). Conversely, activin enhances FSH biosynthesis and secretion and is involved in apoptosis, fibrosis, inflammation, and neurogenesis. Because of its essential role, activin is regarded as a novel drug target (Tsuchida et al., 2009). Myostatin, also referred as growth and differentiation factor 8 (GDF-8), modulates skeletal muscle growth and has been a therapeutic target for disease conditions such as muscular dystrophy, sarcopenia, cashexia, and diabetes mellitus (Walsh and Celeste, 2005). The signaling pathway through TGF-b receptors involves the binding of an extracellular ligand to a type II receptor. The ligand/type II receptor complex phosphorylates a type I receptor via serine/threonine kinase domains of the respective receptors. The signal is further propagated into the cell, initially by phosphorylation of Smad proteins (Attisano et al., 1996; Graham and Peng, 2006; Shi and Massague´, 2003). Both type I and type II receptors are glycoproteins of approximately 55 and 70 kDa, respectively. A distinction between the two types of receptors is based on sequence conservation within the kinase domains and presence of a highly conserved glycine–serine-rich (GS) domain in the cytoplasmic region of type I receptors. Two different activin type II receptors, ActRIIA and ActRIIB, have been characterized. Biochemical studies showed that both ActRIIA and ActRIIB bind activin with high affinity and ALK4 (activin receptor type IB, ActRIB) is the primary type I receptor (Attisano et al., 1992; Mathews and Vale, 1991). In contrast, myostatin binds to ActRIIB more effectively than to ActRIIA (Lee and McPherron, 2001) and ALK5 (TGF-b receptor I, TbRI) is the myostatin type I receptor (Donaldson et al., 1992; Shinozaki et al., 1992). Signaling of activin and myostain is uniquely controlled by intracellular adaptor and scaffolding proteins containing PZD domains (Tsuchida, 2004). A number of TGF-b ligand structures have been determined, revealing a common cysteine knot protein fold (McDonald and Hendrickson, 1993; Sun and Davies, 1995). Furthermore, the structural studies of the extracellular domain of ActRIIB in complex with ligands have been performed (Greenwald et al., 2003, 2004; Thompson et al., 2003). The crystal structures of the cytoplasmic portion of the type I TbRI in different phosphorylation states revealed the key feature of the TbRI activation process (Huse et al., 1999, 2001). The three-dimensional (3D) structure of the cytoplasmic domain of ActRIIB, which contains the catalytic kinase domain, provided insights into the enzyme–substrate complex formation and rational design of selective inhibitors (Han et al., 2007).
Crystal Structure of Human ActRIIB Kinase Domain
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Based on available crystal structures of TbRI and ActRIIB, the conserved and specific features of kinase domains of type I and type II receptors are discussed and the structural analysis will be of help for developing selective inhibitors.
II. Type I Receptor Kinase Domain Structures A. TGF-b RI kinase domain structure Most of the structural studies of type I receptor cytoplasmic domain have been performed using the TbRI receptor. The TbRII, the type II receptor phosphorylates multiple serine and threonine residues in the TTSGSGSG sequence of the cytoplasmic GS domain of the TbRI, leading to its activation (Shi and Massague´, 2003). The GS domain is located upstream of the serine/threonine kinase domain in the cytoplasmic portion of the receptor and serves as an important regulatory domain for TGF-b signaling. In the crystal structure of TbRI cytoplasmic domain complexed with FKBP12, the immunophilin FKBP12 binds to the unphosphorylated GS region and locks the kinase catalytic residues in inactive conformation (Fig. 2.1A; Huse et al., 1999). The tetraphosphorylation of the GS region of TbRI by in vitro protein ligation strategy allowed the GS region from a binding site for an inhibitor into a binding surface for Smad2 substrate. Furthermore, the tetraphosphorylated TbRI is no longer recognized by the inhibitory protein FKBP12 (Huse et al., 2001).
B. ActRIA structure in complex with FKPB12 and dorsomorphin ActRIA (activin receptor type IA, also known as ACVRI) belongs to the bone morphogenetic protein (BMP) receptor family of transmembrane serine/threonine kinases. The structure of the GS and kinase domains of ˚ resolution is ActRIA in complex with FKBP12 and dorsomorphin at 2.35 A available in the Protein Data Bank (PDB code: 3H9R, www.rcsb.org/) (Fig. 2.1B). A comparison of the structures of FKBP12 bound to the TbRI and ActRIA reveals a significant overlap in overall structure including the FBKP-binding site and ATP-binding site with an overall root mean square ˚ for 274 Ca atom pairs. deviation (RMSD) value of 0.58 A A substitution of adenine for guanine at nucleotide 617 replaces a conserved arginine with histidine at residue 206 in the GS domain of ActRIA, which is the primary cause of fibrodysplasia ossificans progressiva (FOP). The FOP is a rare autosomal dominant disorder of skeletal malformations and progressive extraskeletal ossification (Shore et al., 2006). It has been postulated that substitution with histidine creates pH-sensitive
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A
B FKBP12
FKBP12
GS region
Arg206
GS region
Dorsomorphin
b 10 b9
b9
Figure 2.1 Type I receptor kinase domain structures. (A) The structure of TbRI in complex with FKPB12. (B) ActRI kinase domain in complex with FKBP12 and dorsomorphin.
switch within the activation domain of the receptor, leading to ligandindependent activation of ActRIA (Groppe et al., 2007; Fig. 2.1B).
III. 3D Structure of ActRIIB A. Overall structure The crystal structure of unphosphorylated ActRIIB kinase domain from ˚ resolution is very similar to other kinase catalytic domains, human at 2.0 A displaying a bilobal architecture. The smaller N-terminal lobes contain a five-stranded antiparallel b sheet and a single a helix (aC) (Fig. 2.2). The larger C-terminal lobe is mostly a helical and contains the activation loop involved in polypeptide substrate binding. N- and C-terminal lobes are connected by the so-called hinge sequence, which partially defines the binding site for ATP and ATP-competitive kinase inhibitors (Fig. 2.3A). The catalytic domain of the ActRIIB exhibits strong overall sequence similarity with TbRI and ActRIA (39% and 38% identity, respectively). Least-squares Ca superposition of the ActRIIB and TbRI kinase domain
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Crystal Structure of Human ActRIIB Kinase Domain
b1
b1 b4
b4
b5
b5
b3
b3 b2
b2
aC
aC b7
b7 b 10
b9
b 11
b6 b8
aD aE
b 10
b 11
b6 b8
aD
aE
a EF1 a EF2
aF
b9
a EF1 a EF2
aF aJ
aJ aI
aI aG
aG aH
aH
Figure 2.2 Stereoview of the ActRIIB–adenine complex. Secondary structure elements are shown in orange (a helices), cyan (b strands), and gray (loops). The bound adenine is shown as spheres. Activation loop is colored in red.
shows their similarity and yields an overal RMSD value of 1.2 A˚ for 281 Ca atom pairs. When compared with ActRIA kinase domain structure, ˚ ActRIIB catalytic domain overalps with an overall RMSD value of 1.2 A for 269 Ca atom pairs. Differences in the Ca backbones are more pronounced in solvent-exposed loops. The activation loop (residues 339–368), which shows substantial squence variation and flexibility among kinases, clearly distinguishes ActRIIB from TbRI (Figs. 2.2 and 2.3). In both FKBP12-bound TbRI and TbR1-NPC-30345 complexes, the activation loop forms a b hairpin (b9 and b10) that is supported by a one and a half turn extension of the aF helix (Huse et al., 1999, 2001; Fig. 2.1A). However, in the adenine-bound ActRIIB structure, the b hairpin is absent and the equivalent b10 is directly connected to an additional one and a half turn helix (aEF1) (Fig. 2.2). Furthermore, in FKBP-bound ActRI structure, the corresponding activation loop is disordered (Fig. 2.1B).
B. Active site In the ActRIIB–adenine structure, the adenine molecule binds to a considerably different orientation from the adenine moiety of ATP observed in other kinase structures. The adenine inserts into a hydrophobic pocket
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b1
b4
b2
b3
b5
b7 b8
aD
b9
a EF2
aC
b 10
aE
b 11
a EF1
aF
aH
b6
aG
aI
aJ
Figure 2.3 An alignment of the ActRIIB with other type I and type II receptor kinase domains (ActRIA, ActRIB, ActRIIA, TbRI, and TbRII). Secondary structure elements of the ActRIIB are represented by “noodles” (a helices) and “arrows” (b strands). They are colored orange for helices and blue for strands. Identical residues are highlighted in magenta. The conserved residues involved in forming the back pocket are shown as “” below the sequences. The gatekeeper residue is shown as “*” below sequences.
formed by the side chains of Ala215, Leu245, Phe267, Leu328, and Ala338. Instead of N1 nitrogen and amino group, the N3 nitrogen and the protonated N9 of the adenine ring hydrogen bond to the main chain of Ala266 and His268, respectively (Fig. 2.4). The amino group of adenine is solventexposed without interaction with protein. The different binding mode of adenine seen in ActRIIB compared to adenine moeity of ATP binding seen in many other kinases is due to lack of additional intereaction by
35
Crystal Structure of Human ActRIIB Kinase Domain
Lys217
Ala215
Phe267
Phe234 Glu230
Thr265 Leu245
Ala338 Leu328
Gln358
Val369 Leu370
Figure 2.4 Active site of ActRIIB bound to adenine. The hydrogen bonds of adenine (magenta) and Gln358 (pink) in activation loop (gray) are indicated by broken lines. Residues involved in lipophilic pocket for adenine are shown in orange and gatekeeper residue in green.
ribose–phosphate moieity and presence of tautomers with two different protonation states in N9 nitrogen. Both ADP molecule and Mg2þ can be docked into the active site of ActRIIB structure with small changes in glycinerich loop as Ala197 clashes with the ribose moiety of the ADP molecule. The conserved salt bridge between Glu230 from aC helix and Lys217 from b3 strand is not formed in the ActRIIB–adenine structure. Instead, Glu230 forms water-bridged hydrogen bond with Lys217 and a hydrogen bond with the backbone amine of Phe340, which is part of the conserved DFG sequence that marks the N-terminal end of the activation loop. Despite the missing Lys217-Glu230 salt bridge, the adenine binding induces the unphosphorylated activation loop to adopt a conformation similar to that of the fully active form (Fig. 2.4).
C. Comparison to type I/II receptor kinase domain structures Thr265, a conserved gatekeeper residue in ActRIIB, connecting N-terminal domain and the hinge loop (Ala266-Gly271) plays an important structural role by forming a water-bridged hydrogen bond with carbonyl backbone of Leu263 and is involved in forming an additional lipophilic pocket with Phe234, Leu245, and Phe247. The gatekeeper residue in both TbRI and ActRIB is serine residue (Fig. 2.4). In the crystal structure of TbRI complexed with NPC-30345, the size of side chain in the gatekeeper residue played an important role in specificity of NPC-30345 for TbRI (Huse et al., 2001).
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Gln358, the conserved residue in activation loop of ActRIIB and TbRII, is stabilized by hydrogen bonds with carbonyl backbones of Val369 and Leu370 (Fig. 2.4). However, the structurally equivalent residue, Arg372 in TbRI–FKBP12 complex, disrupts ATP-binding site by extending its side chain into the catalytic center to form an ion pair with Asp351, a ligand for an important magnesium ion (Huse et al., 1999) (Fig. 2.3). The ActRIA–FKBP12 complex, similar to TbRI–FKBP12 complex, possesses Arg375 disrupting ATP-binding site by foriming ion pair with Asp354. The Phe234 at the end of the aC helix is involved in forming a back pocket and plays a structural role by hydrophobic interaction with two phenylalanine residues: Phe247 in b4 strand and Phe340 in the conserved DFG motif. In contrast, TbRI bears a large tyrosine residue, Tyr249, which has to move into the back end of the ATP-binding pocket to convert the N-terminal lobe into catallytically active conformation (Huse et al., 2001). The Tyr249 in TbRI crystal structure is involved in hydrophobic interaction with Phe262 and Leu362, and is further stabilized by a hydrogen bond with the carbonyl backbone of Leu260. Similar interaction is observed in the ActRIA–FKPB12 structure, where Tyr252 forms hydrophobic interaction with Phe265 and Leu355 and a hydrogen bond with the carbonyl backbone of Leu263.
IV. Conclusion Structural studies of kinase domains of type I/II receptors have been performed. Small structural differences and plasticity between the ATPbinding sites of these receptor kinases have been successfully exploited to achieve selectivity and potency (Noble and Endicott, 1999; Wang et al., 1998). The crystal structure of human ActRIIB–adenine complex reveals the unphosphorylated activation loop to adopt a conformation similar to that of the fully active form, in spite of the missing Lys217-Glu230 salt bridge. The AcRIIB kinase domain from human represents a distinct typeII receptor serine/threonine kinase subfamily identifiable in part by common features of Thr265 as a gatekeeper residue and back pocket supported by Phe247. The human ActRII kinase domain structure thus provides a basis for a more integrated understanding of substrate recognition and catalysis and will also be of help for developing chemical inhibitors.
REFERENCES Attisano, L., Wrana, J. L., Cheifetz, S., and Massague´, J. (1992). Novel activin receptors: Distinct genes and alternative mRNA splicing generate a repertoire of serine/threonine kinase receptors p97. Cell 68, 97–108.
Crystal Structure of Human ActRIIB Kinase Domain
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Attisano, L., Wrana, J. L., Montalvo, E., and Massague´, J. (1996). Activation of signalling by the activin receptor complex. Mol. Cell. Biol. 16, 1066–1073. Donaldson, C. J., Mathews, L. S., and Vale, W. W. (1992). Molecular cloning and binding properties of the human type II activin receptor. Biochem. Biophys. Res. Commun. 184, 310–316. Graham, H., and Peng, C. (2006). Activin receptor-like kinases: Structure, function and clinical implications. Endocr. Metab. Immune Disord. Drug Targets 6, 45–58. Greenwald, J., Groppe, J., Gray, P., Wiater, E., Kwiatkowski, W., Vale, W., and Choe, S. (2003). The BMP7/ActRII extracellular domain complex provides new insights into the cooperative nature of receptor assembly. Mol. Cell 11, 605–617. Greenwald, J., Vega, M. E., Allendorph, G. P., Fischer, W. H., Vale, W., and Choe, S. (2004). A flexible activin explains the membrane-dependent cooperative assembly of TGF-b family receptors. Mol. Cell 15, 485–489. Groppe, J. C., Shore, E. M., and Kaplan, F. S. (2007). Functional modeling of the ACVR1 (R206H) mutation in FOP. Clin. Orthop. Relat. Res. 462, 87–92. Han, S., Loulakis, P., Griffor, M., and Xie, Z. (2007). Crystal structure of activin receptor type IIB kinase domain from human at 2.0 A˚ resolution. Protein Sci. 16, 2272–2277. Huse, M., Chen, Y.-G., Massague´, J., and Kuriyan, J. (1999). Crystal structure of the cytoplasmic domain of the type I TGFb receptor in complex with FKBP12. Cell 96, 425–436. Huse, M., Muir, T. W., Xu, L., Chen, Y.-G., Kuriyan, J., and Massague´, J. (2001). The TGFb receptor activation process: An inhibitor- to substrate-binding switch. Mol. Cell 8, 671–682. Lee, S. J., and McPherron, A. C. (2001). Regulation of myostatin activity and muscle growth. Proc. Natl. Acad. Sci. USA 98, 9306–9311. Mathews, L. S., and Vale, W. W. (1991). Expression cloning of an activin receptor, a predicted transmembrane kinase. Cell 65, 973–982. McDonald, N. Q., and Hendrickson, W. A. (1993). A structural superfamily of growth factors containing a cystine knot motif. Cell 73, 421–424. McPherron, A. C., Lawler, A. M., and Lee, S. J. (1997). Regulation of skeletal muscle mass in mice by a new TGF-b superfamily member. Nature 387, 83–90. Noble, M. E., and Endicott, J. A. (1999). Chemical inhibitors of cyclin-dependent kinases: Insights into design from X-ray crystallographic studies. Pharmacol. Ther. 82, 269–278. Shi, Y., and Massague´, J. (2003). Mechanisms of TGF-b signaling from cell membrane to the nucleus. Cell 113, 685–700. Shinozaki, H., Ito, I., Hasegawa, Y., Nakamura, K., Igarashi, S., Nakamura, M., Miyamoto, K., Eto, Y., Ibuki, Y., and Minegishi, T. (1992). Cloning and sequencing of a rat type II activin receptor. FEBS Lett. 312, 53–56. Shore, E. M., Xu, M., Feldman, G. J., Fenstermacher, D. A., Cho, T. J., Choi, I. H., Connor, J. M., Delai, P., Glaser, D. L., LeMerrer, M., Morhart, R., Rogers, J. G., et al. (2006). A recurrent mutation in the BMP type I receptor ACVR1 causes inherited and sporadic fibrodysplasia ossificans progressiva. Nat. Genet. 38, 525–527. Sugino, H., and Tsuchida, K. (2000). Activin and follistatin. In “Skeletal Growth Factor,” (E. Canalis, Ed.). Lippincott Williams & Wilkins, Philadelphia, pp. 251–263. Sun, P. D., and Davies, D. R. (1995). The cysteine-knot growth-factor superfamily. Annu. Rev. Biophys. Biomol. Struct. 24, 269–291. Thompson, T. B., Woodruff, T. K., and Jardetzky, T. S. (2003). Structures of an ActRIIB: activin A complex reveal a novel binding mode for TGF-b ligand:receptor interactions. EMBO J. 22, 1555–1566. Tsuchida, K. (2004). Activins, myostatin and related TGF-b family members as novel therapeutic targets for endocrine, metabolic and immune disorders. Curr. Drug Target Immune Endocr. Metabol. Disord. 4, 157–166.
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Tsuchida, K., Nakatani, M., Hitachi, K., Uezumi, A., Sunada, Y., Ageta, H., and Inokuchi, K. (2009). Activin signaling as an emerging target for therapeutic interventions. Cell Commun. Signal. 7, 15–25. Walsh, F. S., and Celeste, A. J. (2005). Myostatin: A modulator of skeletal-muscle stem cells. Biochem. Soc. Trans. 33, 1513–1517. Wang, Z., Canagarajah, B. J., Boehm, J. C., Kassisa, S., Cobb, M. H., Young, P. R., AbdelMeguid, S., Adams, J. L., and Goldsmith, E. J. (1998). Structural basis of inhibitor selectivity in MAP kinases. Structure 6, 1117–1128.
C H A P T E R
T H R E E
Activin/Nodal Signaling and Pluripotency Zhenzhi Chng,* Ludovic Vallier,† and Roger Pedersen† Contents 40 41 44 45
I. Introduction A. The origins of stem cells B. Differences between mouse and human ESCs C. Induced-pluripotent stem cells D. The function of Activin/Nodal/TGFb signaling in stem cell pluripotency II. Conclusion References
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Abstract Maintenance of a pluripotent cell population during mammalian embryogenesis is crucial for the proper generation of extraembryonic and embryonic tissues to ensure intrauterine survival and fetal development. Pluripotent stem cells derived from early stage mammalian embryos are known as “embryonic stem cells.” Such embryo-derived stem cells can proliferate indefinitely in vitro and give rise to derivatives of all three primary germ layers. Their potential for clinical and commercial applications has sparked great excitement within scientific and lay communities. Identification of the signaling pathways controlling stem cell pluripotency and differentiation provides knowledge-based approaches to manipulate stem cells for regenerative medicine. One of the signaling cascades that has been identified in the control of stem cell pluripotency and differentiation is the Activin/Nodal pathway. Here, we describe the differences among pluripotent cell types and discuss the latest findings on the molecular mechanisms involving Activin/Nodal signaling in controlling their pluripotency and differentiation. ß 2011 Elsevier Inc.
* Institute of Medical Biology, Singapore, Singapore Anne McLaren Laboratory for Regenerative Medicine, University of Cambridge, Cambridge, United Kingdom
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Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00003-2
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2011 Elsevier Inc. All rights reserved.
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I. Introduction Embryonic stem cells (ESCs) are pluripotent cells derived from embryos at the blastocyst stage. They possess the dual properties of pluripotency and self-renewal, which means that they are able to proliferate indefinitely in vitro, while maintaining the ability to differentiate into derivatives of all three primary germ layers. Human ESCs (Thomson et al., 1998) offer a potential source of replacement cells to treat diseases such as Parkinson’s (Newman and Bakay, 2008), spinal cord injury (Ronaghi et al., 2010), diabetes mellitus (Heit and Kim, 2004), and cardiomyopathies ( Janssens, 2007). In addition to their potential for cell therapies, the embryonic origin of ESCs confers upon them the ability to recapitulate the early events occurring in the developing embryo. Therefore, understanding the molecular mechanisms controlling pluripotency and differentiation of ESCs could provide further insights into early embryonic development, an area of study that has been limited especially in humans due to the difficulty in obtaining experimental material and the ethical concerns involving human embryo research. Despite their common origin and similar properties of self-renewal and pluripotency, human ESCs and mouse ESCs do not rely on the same signaling pathways to maintain their pluripotent status. While mouse ESCs depend on leukemia inhibitory factor (LIF) and bone morphogenetic protein (BMP) signaling to remain pluripotent, human ESCs depend on Activin/Nodal signaling and fibroblast growth factor-2 (FGF2) signaling to maintain their pluripotent status (Beattie et al., 2005; Besser, 2004; Greber et al., 2007; James et al., 2005; Ludwig et al., 2006a,b; Vallier et al., 2004, 2005; Xiao et al., 2006; Yao et al., 2006). The isolation of pluripotent cells derived from the late epiblast layer of postimplantation mouse, rat, and pig embryos (epiblast stem cells [EpiSCs]) has led to the hypothesis that human ESCs represent the epiblast stage of human embryonic development rather than the blastocyst stage (Alberio et al., 2010; Brons et al., 2007; Tesar et al., 2007), and that Activin/Nodal signaling plays the key role in maintaining pluripotency of cells at this stage of development. Later studies have also shown that induced-pluripotent stem cells (iPSCs), which are generated from somatic cells that have been genetically reprogrammed to an ESC-like state (Takahashi et al., 2007), also depend on Activin/Nodal signaling to remain pluripotent (Vallier et al., 2009b). This chapter provides a brief explanation of the different types of pluripotent stem cells that depend on Activin/Nodal signaling, and describes in detail the function of Activin/ Nodal signaling pathway in maintaining self-renewal and pluripotency in these cell types.
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A. The origins of stem cells 1. Locations of pluripotent cells during embryonic development In mammals, fertilization occurs in the oviduct, a tube linking the ovary and the uterus. The first cleavage of the zygote occurs about 24 h later as the embryo is swept toward the uterus by cilia and peristaltic contractions occurring in the oviduct. During transit to the uterus, the zona pellucida surrounding the embryo prevents it from prematurely adhering to the oviduct. In humans, around 4 days after fertilization, the embryo undergoes compaction, forming the morula, a compact ball of cells. The morula consists of a small group of internal cells surrounded by a larger group of external cells. Additional cells are recruited to the inner group as the embryo progresses from the morula to the blastocyst stage, a hollow ball of cells whose inner cells constitute the inner cell mass (ICM), from which the embryo proper develops, and whose external cells (the trophoblast) become the placenta. This early segregation of the ICM and trophoblast cells represents the first differentiation event in mammalian development. The embryo acquires the blastocyst cavity when its cells retain fluid transported across its developing trophoblast cell layer. The ICM then occupies the embryonic pole of the preimplantation blastocyst. Around the end of the fifth day, the embryo “hatches” from the zona pellucida and implants into the uterine wall. Extensive extraembryonic tissues continue to form upon implantation. Particularly, the human syncytiotrophoblast cells actively invade the uterine wall to increase the surface area available for nutrient exchange between the mother and the fetus (Gilbert, 2003). During the peri-implantation stages (Fig. 3.1), a population of pluripotent cells remains sequestered from inductive influences that would result in their precocious differentiation. These undifferentiated, pluripotent cells are found in the ICM of the preimplantation blastocyst and in the innermost (early epiblast) layer of the peri-implantation, late blastocyst ICM. The early epiblast is formed after hypoblast cells delaminate from the ICM to line the blastocoel cavity and contains cells that can differentiate into the three primary germ layers that form the fetus, as well as the extraembryonic mesoderm of the yolk sac, amnion, and chorion. The early epiblast cell layer is regarded as the source of mouse ESCs (Evans and Kaufman, 1981; Martin, 1981) on the basis of the requirement of gp130 signaling for epiblast survival at this stage (Nichols et al., 2001). With formation of the proamniotic cavity early on the sixth day of gestation (E5.0þ) of mouse embryos, the growth pattern of early epiblast changes, marking its transition into a simple epithelium with noncoherent clonal growth (cell mixing), accompanied and likely caused by interkinetic nuclear migration (Gardner and Cockroft, 1998; Lawson et al., 1991). Culture of late epiblast layers yielded EpiSCs (Brons et al., 2007; Tesar et al., 2007), which can thus be regarded as originating from the epithelial stage of pluripotent embryonic cells, just before the onset of gastrulation.
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Embryonic pole ICM Trophectoderm Blastocoel Fertilization Preimplantation blastocyst Syncytiotrophoblast Early epiblast
Hypoblast Cytotrophoblast
Postimplantation blastocyst
Amnionic ectoderm
Late epiblast
Amniotic cavity Extraembryonic mesoderm Hypoblast Syncytiotrophoblast
Yolk sac
Implanted embryo
Ectoderm Mesoderm Endoderm Gastrulation
Figure 3.1 Location of pluripotent cells in the early human embryo. In the early stages of mammalian development, extraembryonic tissues differentiate in the outer layers of the conceptus [depicted as trophoblast (gray) or hypoblast (pink)] before, during, and after implantation in the uterine wall. By contrast, the inner/innermost layers of the conceptus remain pluripotent, either as ICM, early epiblast or late epiblast cells (depicted in green). A population of pluripotent cells is thus maintained through gastrulation, when they differentiate into the primary germ layers (ectoderm, mesoderm, and endoderm). ICM: inner cell mass (Reynolds et al., 2009).
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2. Derivation of ESCs Although ESCs research, particularly human ESCs research, is perceived as a phenomenon that emerged in the past decade, the developments in this field are deeply rooted in research and accomplishments that date back to the 1950s. In 1954, Stevens and Little first described an incidence of spontaneous testicular teratomas, which were histologically similar to typical teratomas but were malignant, in the inbred 129/terSv mouse strain (Stevens and Little, 1954). This discovery instigated a series of systematic experimental studies in the 1950s and 1960s on teratomas and teratocarcinomas. A crucial experiment was the demonstration that teratocarcinomas consist of a unique type of stem cells (later known as embryonal carcinoma cells) that have the ability to differentiate into adult cell types and are able to grow indefinitely when injected intraperitoneally (Kleinsmith and Pierce, 1964). In 1970, the discovery that mouse embryos grafted to extrauterine sites give rise to retransplantable teratocarcinomas, introduced the concept that pluripotent stem cells of early mouse embryos and teratocarcinomas might be highly similar (Solter et al., 1970; Stevens, 1970). At that time, Gardner had shown that it was possible to transplant cells between the ICM of different blastocysts, and that these cells would contribute to a chimeric embryo, with cells from the transplant contributing to all germ layers of the adult (Gardner, 1968; Moustafa and Brinster, 1972). These findings demonstrated that the pluripotent cells of the embryo could be found in the ICM, and instigated a series of attempts to isolate and culture ICM cells ex vivo. The first mouse ESCs (mESCs) were derived from mouse ICM in 1981 by Evans and Kaufman (Evans and Kaufman, 1981; Martin, 1981), and this was confirmed by Martin (1981). Despite the difference in derivation techniques, the use of mitotically immortalized mouse embryonic fibroblasts (MEFs) feeder layer was crucial for the culture of mESCs. mESCs remain pluripotent when grown on MEFs, but differentiate in their absence. In mouse chimeras made with intact embryos, mESCs contribute to a wide range of adult tissues and are also capable of germline transmission, thus providing a powerful tool for introducing specific genetic changes into the mouse germline. Due to the difficulties in obtaining suitable human embryonic material and the ethical issues that accompany human embryo research, the derivation of human ESCs (hESCs) lagged significantly behind their murine counterparts. In 1995, Thomson isolated the first primate ESC line from rhesus monkey blastocyst (Thomson and Marshall, 1998) and in 1998, successfully isolated the first hESC line from the ICM of fresh or frozen cleavage-stage human embryos produced by in vitro fertilization (Thomson et al., 1998). These hESC lines have normal karyotypes, express high levels of telomerase activity, and express cell-surface markers that characterize primate ESCs including stage-specific embryonic antigen (SSEA)-3, SSEA-4, TRA-l-60, TRA-1-81, and alkaline phosphatase.
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After undifferentiated proliferation in vitro for 4–5 months, these cells maintained the developmental potential to form trophoblast and derivatives of all three germ layers, including neural epithelium, embryonic ganglia, and stratified squamous epithelium (ectoderm); cartilage, bone, smooth muscle, and striated muscle (mesoderm); and gut epithelium (endoderm). This demonstration of pluripotency for human ICM-derived stem cells sparked interest in potential applications of hESCs for human therapies and thereby accelerated research on mammalian stem cells.
B. Differences between mouse and human ESCs The initial conditions used for derivation and culture of ESCs were identical for mouse and human and consisted of mouse fetal (embryonic) fibroblasts (MEFs) and fetal calf serum, thus comprising an essentially undefined growth factor milieu (Evans and Kaufman, 1981; Martin, 1981; Thomson et al., 1998). It was therefore unclear which components of the culture conditions were responsible for maintenance of pluripotency. In studies on mESCs, conditioned media from buffalo rat liver cells were found to be an effective substitute for MEFs to block differentiation (Smith and Hooper, 1987). Analysis of the components of the conditioned medium revealed that Leukemia inhibitory factor (LIF), a cytokine known to induce macrophage differentiation of myeloid leukemic cells (Gearing et al., 1987), could substitute for MEFs in the maintenance of mESCs in a pluripotent state in vitro (Nichols et al., 1990; Smith et al., 1988; Williams et al., 1988). It was later found that LIF maintains mESC pluripotency via activation of STAT3 (Boeuf et al., 1997; Niwa et al., 1998). While LIF is sufficient to maintain mESC pluripotency in the absence of MEFs, it does not substitute for serum, without which mESCs undergo neural differentiation. Ying et al. discovered that the induction of Id proteins by BMP signaling suppresses neural differentiation and sustains mESC self-renewal in collaboration with STAT3 (Ying et al., 2003). Thus, a chemically defined feeder-free and serum-free culture of mESCs was established, based on combined use of LIF and BMP. Despite their apparently common developmental origin and shared properties of self-renewal and pluripotency, hESCs and mESCs rely on different signaling pathways to maintain their pluripotent status. LIF and BMP signaling do not maintain hESC pluripotency (Daheron et al., 2004; Humphrey et al., 2004). Instead, Activin/Nodal and FGF2 signaling promote hESC pluripotency (Amit et al., 2004; Beattie et al., 2005; Besser, 2004; Greber et al., 2007; James et al., 2005; Levenstein et al., 2006; Ludwig et al., 2006a; Vallier et al., 2005; Xiao et al., 2006; Yao et al., 2006). Instead, BMP drives hESC differentiation towards extraembryonic lineages (Hayashi et al., 2009; Vallier et al., 2009c; Xu, 2006). In addition, unlike mESCs, which form compact colonies and propagates as single cells, hESCs form flat colonies and cannot grow efficiently as single cells (Amit et al., 2000).
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While mESCs cannot differentiate into trophectoderm or primitive endoderm (Beddington and Robertson, 1989), hESCs differentiate into extraembryonic tissues when treated with BMP4. The explanation for these differences between mESCs and hESCs has been unclear, until two groups independently derived pluripotent stem cells from the epiblast layer of postimplantation murine blastocysts (Brons et al., 2007; Tesar et al., 2007). There are several striking similarities between mEpiSCs and hESCs: they depend on Activin/Nodal and FGF signaling to maintain pluripotency; they differentiate into cells expressing trophectoderm markers when induced with BMP4; they have limited capacity to survive as single cells; and they are unable to efficiently form chimeras when reintroduced into the mouse blastocyst. This suggests that the differences between mESCs and hESCs are due to the different developmental stages that these two cell types represent, rather than being attributable to a species difference. The dependence on LIF and BMP signaling by mESCs replicates the response of early epiblast cells to the signaling environment experienced by the mouse ICM during diapause1 (Nichols et al., 2001). The dependence on Activin/Nodal and FGF signaling by hESCs and mEpiSCs may similarly replicate the response to the signaling environment shown by the mouse late epiblast during postimplantation stages. Human ESCs are derived by culturing embryos from the blastocyst stage. The dependence of hESCs on similar signaling pathways as mEpiSCs implies that the cultured human ICM cells progress in vitro from the ICM stage to the late epiblast stage during the process of hESC derivation (Brons et al., 2007).
C. Induced-pluripotent stem cells While hESC lines from normal embryos are useful in vitro models for studying mechanisms controlling early human development (Vallier and Pedersen, 2005), stem cell lines from patients with genetic or acquired diseases would provide powerful tools for research into disease progression, and lay a foundation for producing autologous cell therapies that would avoid immune rejection and correction of gene defects prior to tissue reconstitution (Park et al., 2008a). With this goal in mind, hESC lines from embryos shown to carry genetic diseases by virtue of preimplantation genetic diagnosis have been isolated (Verlinsky et al., 2005). However, those cell lines only represent common, monogenic conditions. An alternative strategy for producing autologous, patient-derived pluripotent stem cells is to “dedifferentiate” a patient’s somatic cell by somatic cell nuclear transfer, cell fusion, induction of pluripotency by ectopic gene expression, or direct 1
Diapause is an implantational delay that results in the blastocyst entering a state of metabolic and proliferative quiescence. During murine pregnancy, LIF is also required to induce the uterus to become receptive to the blastocyst to allow implantation of occur (Hondo and Stewart, 2005).
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reprogramming (Gurdon and Melton, 2008). A promising approach for modeling human diseases has arisen from Yamanaka’s success in reprogramming somatic cells by introducing a set of transcription factors linked to pluripotency (c-Myc, Klf4, Oct4, and Sox2) to produce Induced-pluripotent stem cells (iPSCs) (Takahashi and Yamanaka, 2006; Takahashi et al., 2007). The advent of this iPSC technology has instigated great enthusiasm in the stem cell community worldwide to explore this technology for applications in disease modeling, patient-disease-specific drug discovery, and gene correction therapy (Lowry et al., 2008; Park et al., 2008b; Takahashi et al., 2007; Yu et al., 2007). It has been shown that similar signaling pathways control early cell fate decisions of human iPSCs and hESCs (Vallier et al., 2009b). Therefore, current knowledge on direct differentiation of hESCs could be adapted for human iPSCs. However, the use of iPSCs in the clinic still faces many hurdles. Current methods for reprogramming require infecting the somatic cells with multiple viral vectors (Lowry et al., 2008; Park et al., 2008b; Takahashi et al., 2007; Yu et al., 2007). Replacing the use of viral vectors by small molecules (Zhou et al., 2009) or plasmid vectors (Okita et al., 2010) would reduce risk of random integration by the viral vectors which may cause mutation. In addition, other underlying problems of iPSCs such as their elusive origin and the risk of tumorigenesis (Ou et al., 2010) will have to be addressed before iPSCs can be fully utilized for transplantation. Insight into these issues may be gained by understanding the molecular mechanisms by which pluripotency is maintained and differentiation is induced in hESCs, EpiSCs, and human iPSCs. The following sections thus describe in detail how Activin/Nodal signaling is thought to function in maintaining self-renewal and pluripotency of hESCs, human iPSCs, and mEpiSCs in vitro, and also in the early mouse embryo.
D. The function of Activin/Nodal/TGFb signaling in stem cell pluripotency 1. Activin/Nodal/TGFb pathway Activin and Nodal belong to the transforming growth factor b (TGFb) superfamily of growth factors, which consists of more than 40 ligands including TGFbs, inhibins, myostatins, BMPs, growth/differentiation factors (GDFs), and the anti-Mu¨llerian hormone. Expression of these growth factors can be detected in most embryonic tissues and their adult derivatives. In agreement with their ubiquitous expression, TGFb ligands, receptors, and their downstream cascades have pleiotropic functions, which include the control of proliferation, differentiation, apoptosis, and cell adhesion. TGFb superfamily ligands bind to cell-surface serine/threonine kinase receptors known as TGFb type I and type II receptors. The type II receptor phosphorylates and activates the type I receptor, also known as an Activin receptor-like kinase (ALK). TGFb and Activin have a high affinity for the
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type II receptor but do not bind to the type I receptor in the absence of type II receptor, whereas BMP has higher affinity for the type I receptor than for type II receptor. Being downstream in the Activin/Nodal/TGFb cascade from the type II receptor, the type I receptor determines the specificity of intracellular signals (Valdimarsdottir and Mummery, 2005). TGFb signals are propagated by intracellular mediators, the receptor-activated SMADs (R-SMADs), through two main branches: (1) the BMP branch, which transduces through R-SMADs 1, 5, and 8 via the type I receptors ALK1, ALK2, ALK3, and ALK6 and (2) the TGFb/Activin/Nodal branch, which involves the activation of R-SMADs 2 and 3 via ALK4, ALK5, and ALK7. Upon activation by phosphorylation, R-SMADs associate with SMAD4, the common SMAD of both the BMP and TGFb/Activin/Nodal pathways. The R-SMAD–SMAD4 complex translocates from the cytoplasm into the nucleus, and together with other transcription cofactors, regulates target gene expression (Attisano and Lee-Hoeflich, 2001). There are two inhibitory SMADs: SMAD6, which inhibits SMADs 1 and 5 (Nakayama et al., 1998) and SMAD7, which inhibits both TGFb/Activin/Nodal and BMP pathways (Casellas and Brivanlou, 1998; Fig. 3.2). Although Nodal and Activin activate the same SMAD2/3 signaling pathway, Nodal uniquely requires Cripto as a cofactor to activate the ALK4/ActRIIb receptors, and only Nodal is inhibited by Lefty and Cerberus (Schier, 2003). TGFb also activates the SMAD2/3 pathway using type I receptor ALK5 and type II receptor TGFbRII, and it can also phosphorylate SMAD1 through ALK1 (Wrighton et al., 2009; Fig. 3.2). SMAD proteins are molecules of 42–60 kDa in size and are characterized by a three-domain structure. The N-terminal MH1 and C-terminal MH2 domains flank a variable proline-rich linker region (Kretzschmar and Massague, 1998). In the absence of signaling, SMADs are kept in a latent conformation through intramolecular interaction between the MH1 and MH2 domains. Activation by phosphorylation reportedly disrupts this autoinhibition, making the MH1 and MH2 domains available for interaction with DNA and other proteins, respectively. SMAD4 and the MH1 domain of activated SMAD3 have been reported to bind directly to DNA, but SMADs in general have low DNA-binding affinity and specificity. However, they are able to achieve highly specific regulation of target promoters through interaction with other cofactors and transcription factors (Attisano and Lee-Hoeflich, 2001; Attisano and Wrana, 2002). To date, more than 50 SMAD partners have been characterized. These include transcriptional coactivators (such as p300/CPB and p/CAF), transcriptional corepressors (such as Ski and SnoN), and many other transcription factors (such as FOXH1, SIP1, and Evi-1) and E3 ubiquitin ligases (such as Ectoderminm and Smurf1/2) (Miyazono et al., 2001). The interaction of SMADs with a large number of proteins partly explains the diversity of mechanisms controlled by TGFb superfamily.
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Ligands
Nodal
TGFb
Activin
BMP
Cripto Lefty/ Cerberus
BMPRII
ActRII Type II receptors
TGFb RII
ActRII
ActRIIB
ActRIIB
ALK2 Type I receptors
ALK4
ALK5
ALK1 ALK3
ALK7
ALK6
R-SMADS
SMAD2
SMAD1
SMAD3
SMAD5 SMAD8
SMAD4
Figure 3.2 Schematic of TGFb signaling through receptors and SMAD cascades. TGFb ligands bind to cell-surface serine/threonine kinase receptors known as TGFb type I and type II receptors. The type II receptor phosphorylates and activates the type I receptor, also known as an Activin receptor-like kinase (ALK). TGFb signals are propagated by intracellular mediators, the receptor-activated SMADs (R-SMADs), through two main branches: (1) the BMP branch, which transduces through R-SMADs 1, 5, and 8 via the type I receptors ALK1, ALK2, ALK3, and ALK6 and (2) the TGFb/Activin/Nodal branch, which involves the activation of R-SMADs 2 and 3 via ALK4, ALK5, and ALK7. Upon activation by phosphorylation, R-SMADs associate with SMAD4, the common SMAD of both the BMP and TGFb/Activin/ Nodal pathways.
2. The function of TGFb signaling in early development of the mouse embryo TGFb superfamily signaling components are expressed ubiquitously in most embryonic tissues and their adult derivatives, and are responsible for diverse cell fate decisions during embryogenesis including the control of proliferation, differentiation, apoptosis, and cell adhesion. Of the large family of ligands, only BMPs, GDFs, and Nodal appear to be necessary for early
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mouse development (reviewed in Reynolds et al., 2009). For example, BMP signaling is required for germ cell specification, proliferation of the epiblast, and differentiation of extraembryonic tissues (Lawson et al., 1999; Mishina et al., 1995; Winnier et al., 1995). The function of GDF3 is less well understood. It has been shown to block BMP signaling and is expressed in pluripotent cells and in the node during gastrulation in a pattern consistent with BMP inhibition (Levine and Brivanlou, 2006). Embryos mutant for GDF3 failed to gastrulate (Chen et al., 2006), suggesting that GDF3 may play a role in pluripotency and early cell fate decisions. However, it has also been shown that high doses of GDF3 activate Nodal signaling (Levine et al., 2009). It is therefore unclear whether GDF3 maintains pluripotency directly or rather acts indirectly through the activation of Nodal transcription. Activin was first purified from gonadal fluids based on its ability to activate the release of follicle-stimulating hormone from cultured pituitary gonadotropes (Ling et al., 1986; Vale et al., 1986). Most studies carried out on the biological function of Activin have focused on its role in reproduction, such as gonadal sex determination (Xia and Schneyer, 2009), follicle development, luteolysis, and testicular function. The first evidence of a role of Activin during early embryo development was suggested by Jim Smith and coworkers, who identified the most potent mesoderm-inducing factor produced by Xenopus XTC cell line as the homologue of mammalian Activin A (Smith et al., 1990). However, Matzuk and coworkers showed that Activin A-deficient mice develop to term but die within 24 h of birth (Matzuk et al., 1995), lacking whiskers and lower incisors and with defects in their secondary palates, including cleft palate. This suggests that while Activin is able to evoke developmental responses in embryonic tissues, Nodal or other TGFb ligands are more likely to be the native factor(s) operating in the embryo. Nodal is initially expressed throughout the mouse epiblast just after implantation and is involved in the specification of the anterior–posterior axis before gastrulation by controlling the development of the anterior visceral endoderm (Robertson et al., 2003; Schier, 2003; Yamamoto et al., 2004). Nodal then becomes highly localized in the posterior part of the embryo where the primitive streak has formed, where it functions to induce the expression of posterior genes such as Wnt3 and Brachyury (Brennan et al., 2001). In the absence of Nodal, mouse embryos arrest at early gastrulation and contain little or no embryonic mesoderm (Conlon et al., 1991, 1994), suggesting that Nodal is required for mesendoderm differentiation. The function of Nodal in mesoderm and endoderm development was also confirmed by gain- and loss-of-function studies carried out in Xenopus and fish embryos ( Jones et al., 1995; Rebagliati et al., 1998). Interestingly, Nodal mutants also suffer from reduced epiblast cell population size and lose expression of the pluripotency marker OCT4 (Varlet et al., 1997), suggesting that the pregastrulation arrest of Nodal-deficient embryos could reflect an impaired pluripotency. This hypothesis was reinforced by studies
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showing that Nodal is necessary at pregastrula stages to maintain the expression of pluripotency markers in the epiblast. Importantly, absence of Nodal results in ectopic expression of neuroectoderm markers (Camus et al., 2006; Mesnard et al., 2006). Taken together, these observations suggest that Nodal functions initially to maintain pluripotency of the epiblast prior to gastrulation, and then acts subsequently in the posterior part of the embryo to control primitive streak formation. 3. The function of TGFb signaling in maintaining stem cell pluripotency Many signaling pathways, including TGFb, Wnt, and Insulin-like growth factor, have been implicated in the maintenance of hESC pluripotency (Reynolds et al., 2009). Feeder-free cultures of hESCs invariably require a source of TGFb superfamily ligands (Amit et al., 2004) and a large number of gene expression profiling studies indicate a function for TGFb signaling in hESCs (Brandenberger et al., 2004a,b; Calhoun et al., 2004; Miura et al., 2004; Rosler et al., 2004). Particularly, the activation of the Activin/Nodal pathway is most commonly described as being sufficient to maintain longterm culture of hESCs in the pluripotent state (Beattie et al., 2005; Besser, 2004; James et al., 2005; Vallier et al., 2005). Although Activin and Nodal have distinct functions in vivo, they activate the same pathway. Addition of either Activin A or recombinant Nodal to cell culture media has the same effects of maintaining stem cell pluripotency. A much larger dose of Nodal is required to achieve the same effects of Activin A, perhaps because Nodal requires cofactors (Cripto and Cryptic) to activate the signaling pathway, or because Nodal is only inefficiently converted from its “pro” to active form. Therefore, Activin is the more commonly used supplement for ESC culture. For simplicity, we use the term “Activin/Nodal signaling” to mean the pathway that utilizes SMAD2 and SMAD3 for signal transduction that could be activated by either of those ligands. Human ESCs, human iPSCs, and mouse EpiSCs cultured in a chemically defined condition devoid of serum or growth factors spontaneously differentiate toward the neuroectoderm lineage (Chng et al. 2010; Vallier et al., 2004, 2009b,c). Activation of Activin/Nodal signaling, either by exogenous administration of Nodal or Activin A or by constitutive overexpression of the Nodal transgene, maintains stem cell pluripotency by blocking the spontaneous neuroectoderm differentiation (Vallier et al., 2004, 2005). Similarly, inhibition of Nodal by overexpression of Lefty increases neuroectoderm differentiation (Smith et al., 2008), and blockade of Activin/Nodal signaling by the pharmacological inhibitor SB4315452 induces differentiation of hESCs grown on feeders, in serum or in chemically defined conditions (Chng et al. 2010; James et al., 2005). Together, these studies support previous in vivo studies showing that Nodal-null epiblast differentiates precociously into neuroectoderm (Camus et al.,
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2006) and suggest an analogous role for Nodal in human and mouse—that is maintaining pluripotency by inhibiting neuroectoderm differentiation and preventing the emergence of anterior and neural cell fates before gastrulation in the late epiblast. Further investigation into the molecular mechanisms revealed that Activin/Nodal and FGF signals synergize to inhibit BMP signaling, thereby sustaining expression of pluripotency genes (such as NANOG, OCT4, and SOX2) and promoting long-term undifferentiated proliferation of hESCs (Xu et al., 2008). In addition, SMAD2/3 (intracellular effectors of Activin/ Nodal) directly bind the promoter NANOG to activate its expression (Vallier et al., 2009a; Xu et al., 2008), while SMAD1 (downstream of BMP signaling) binds the NANOG promoter to downregulate its expression to cause ESC differentiation (Xu et al., 2008). It was also shown that SMAD2/3 cooperates with NANOG, OCT4, and SOX2 to regulate the expression of neuroectoderm-promoting gene Smad-interacting protein 1 (SIP1) (Chng et al., 2010), thus blocking the spontaneous neuroectoderm differentiation of hESCs while maintaining pluripotency. These studies illustrate how the extrinsically driven Activin/Nodal signaling cascade intersects with the intrinsic transcriptional networks to maintain stem cell pluripotency and block neuroectoderm differentiation. However, high doses of Activin in combination with serum or BMP induce differentiation of hESCs into mesoderm and endoderm (D’Amour et al., 2005; McLean et al., 2007; Vallier et al., 2009c), confirming that Activin/Nodal signaling also has a function in hESC differentiation. However, a high dose of Activin alone is not sufficient to cause differentiation of hESCs grown in a chemically defined medium devoid of serum (Vallier et al., 2005, 2009c). This observation suggests that other factors are necessary for Activin/Nodal signaling to effectively induce mesendoderm differentiation, and that Activin/Nodal’s function in maintaining pluripotency and differentiation involves differential recruitment of cofactors for the respective functions. Further investigations on the roles of specific components of the Activin/Nodal signaling pathway (i.e., Smads and their binding partners) in hESCs and during mesoderm and endoderm differentiation, and identification of the target genes controlled by SMAD2/3 transcriptional complexes will provide greater information on the nature of the mechanisms involved in this dual function of the Activin/Nodal signaling cascade.
II. Conclusion Human ESCs, mouse EpiSCs, and human iPSCs offer useful tools for understanding the basic biology of human development and disease progression. These pluripotent cell types are also potentially useful as
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sources of replacement cells for regenerative medicine. Importantly, the stem cell technology can also be used commercially as a platform for drug screening for identification of novel therapeutics. Understanding the signals controlling stem cell pluripotency and differentiation provides important information on the biology of these intriguing cell types, and enables the discovery of ways to differentiate stem cells into clinically and commercially useful cell types. Activin/Nodal signaling is key in maintaining pluripotency of the class of stem cells that includes hESCs, EpiSCs, and hIPSCs. Through the activation of SMAD2/3, Activin/Nodal cooperates with the pluripotency transcriptional circuitry (NANOG, OCT4, and SOX2) to maintain pluripotency and block the spontaneous neuroectoderm differentiation. However, Activin/Nodal signaling is also known to be an inducer of mesendoderm differentiation. These diverse functions of Activin/Nodal signaling can be attributed to the ability of SMADs to interact with a plethora of other intracellular and intranuclear factors. Further investigations into these various SMAD-interacting factors at different stages of stem cell differentiation will provide greater information on the precise role of Activin/Nodal signaling in stem cell development. In addition, understanding the cross talk between Activin/Nodal signaling and other pathways presents (BMP, Wnt, FGF, etc.) is a difficult but important task to enable the establishment of better protocols for in vitro stem cell differentiation and will confer a deeper understanding of how cell fate decisions are made, particularly in human development.
REFERENCES Alberio, R., Croxall, N., and Allegrucci, C. (2010). Pig epiblast stem cells depend on activin/nodal signaling for pluripotency and self renewal. Stem Cells Dev. 19(10), 1627–1636. Amit, M., Carpenter, M. K., Inokuma, M. S., Chiu, C. P., Harris, C. P., Waknitz, M. A., Itskovitz-Eldor, J., and Thomson, J. A. (2000). Clonally derived human embryonic stem cell lines maintain pluripotency and proliferative potential for prolonged periods of culture. Dev. Biol. 227, 271–278. Amit, M., Shariki, C., Margulets, V., and Itskovitz-Eldor, J. (2004). Feeder layer- and serum-free culture of human embryonic stem cells. Biol. Reprod. 70, 837–845. Attisano, L., and Lee-Hoeflich, S. T. (2001). The Smads. Genome Biol. 2(8), reviews 3010.1– 3010.8. Attisano, L., and Wrana, J. L. (2002). Signal transduction by the TGF-beta superfamily. Science 296, 1646–1647. Beattie, G. M., Lopez, A. D., Bucay, N., Hinton, A., Firpo, M. T., King, C. C., and Hayek, A. (2005). Activin A maintains pluripotency of human embryonic stem cells in the absence of feeder layers. Stem Cells 23, 489–495. Beddington, R. S., and Robertson, E. J. (1989). An assessment of the developmental potential of embryonic stem cells in the midgestation mouse embryo. Development 105, 733–737.
Activin/Nodal Signaling and Pluripotency
53
Besser, D. (2004). Expression of nodal, lefty-a, and lefty-B in undifferentiated human embryonic stem cells requires activation of Smad2/3. J. Biol. Chem. 279, 45076–45084. Boeuf, H., Hauss, C., Graeve, F. D., Baran, N., and Kedinger, C. (1997). Leukemia inhibitory factor-dependent transcriptional activation in embryonic stem cells. J. Cell Biol. 138, 1207–1217. Brandenberger, R., Khrebtukova, I., Thies, R. S., Miura, T., Jingli, C., Puri, R., Vasicek, T., Lebkowski, J., and Rao, M. (2004a). MPSS profiling of human embryonic stem cells. BMC Dev. Biol. 4, 10. Brandenberger, R., Wei, H., Zhang, S., Lei, S., Murage, J., Fisk, G. J., Li, Y., Xu, C., Fang, R., Guegler, K., et al. (2004b). Transcriptome characterization elucidates signaling networks that control human ES cell growth and differentiation. Nat. Biotechnol. 22, 707–716. Brennan, J., Lu, C. C., Norris, D. P., Rodriguez, T. A., Beddington, R. S., and Robertson, E. J. (2001). Nodal signalling in the epiblast patterns the early mouse embryo. Nature 411, 965–969. Brons, I. G., Smithers, L. E., Trotter, M. W., Rugg-Gunn, P., Sun, B., Chuva de Sousa Lopes, S. M., Howlett, S. K., Clarkson, A., Ahrlund-Richter, L., Pedersen, R. A., et al. (2007). Derivation of pluripotent epiblast stem cells from mammalian embryos. Nature 448, 191–195. Calhoun, J. D., Rao, R. R., Warrenfeltz, S., Rekaya, R., Dalton, S., McDonald, J., and Stice, S. L. (2004). Transcriptional profiling of initial differentiation events in human embryonic stem cells. Biochem. Biophys. Res. Commun. 323, 453–464. Camus, A., Perea-Gomez, A., Moreau, A., and Collignon, J. (2006). Absence of Nodal signaling promotes precocious neural differentiation in the mouse embryo. Dev. Biol. 295, 743–755. Casellas, R., and Brivanlou, A. H. (1998). Xenopus Smad7 inhibits both the activin and BMP pathways and acts as a neural inducer. Dev. Biol. 198, 1–12. Chen, C., Ware, S. M., Sato, A., Houston-Hawkins, D. E., Habas, R., Matzuk, M. M., Shen, M. M., and Brown, C. W. (2006). The Vg1-related protein Gdf3 acts in a Nodal signaling pathway in the pre-gastrulation mouse embryo. Development 133, 319–329. Chng, Z., Teo, A., Pedersen, R. A., and Vallier, L. (2010). SIP1 mediates cell-fate decisions between neuroectoderm and mesendoderm in human pluripotent stem cells. Cell Stem Cell 6, 59–70. Conlon, F. L., Barth, K. S., and Robertson, E. J. (1991). A novel retrovirally induced embryonic lethal mutation in the mouse: Assessment of the developmental fate of embryonic stem cells homozygous for the 413.d proviral integration. Development 111, 969–981. Conlon, F. L., Lyons, K. M., Takaesu, N., Barth, K. S., Kispert, A., Herrmann, B., and Robertson, E. J. (1994). A primary requirement for nodal in the formation and maintenance of the primitive streak in the mouse. Development 120, 1919–1928. Daheron, L., Opitz, S. L., Zaehres, H., Lensch, M. W., Andrews, P. W., Itskovitz-Eldor, J., and Daley, G. Q. (2004). LIF/STAT3 signaling fails to maintain self-renewal of human embryonic stem cells. Stem Cells 22, 770–778. D’Amour, K. A., Agulnick, A. D., Eliazer, S., Kelly, O. G., Kroon, E., and Baetge, E. E. (2005). Efficient differentiation of human embryonic stem cells to definitive endoderm. Nat. Biotechnol. 23, 1534–1541. Evans, M. J., and Kaufman, M. H. (1981). Establishment in culture of pluripotential cells from mouse embryos. Nature 292, 154–156. Gardner, R. L. (1968). Mouse chimeras obtained by the injection of cells into the blastocyst. Nature 220, 596–597. Gardner, R. L., and Cockroft, D. L. (1998). Complete dissipation of coherent clonal growth occurs before gastrulation in mouse epiblast. Development 125, 2397–2402.
54
Zhenzhi Chng et al.
Gearing, D. P., Gough, N. M., King, J. A., Hilton, D. J., Nicola, N. A., Simpson, R. J., Nice, E. C., Kelso, A., and Metcalf, D. (1987). Molecular cloning and expression of cDNA encoding a murine myeloid leukaemia inhibitory factor (LIF). EMBO J. 6, 3995–4002. Gilbert, S. F. (2003). Developmental Biology seventh ed. Sinauer Associates, Sunderland, MA, USA. Greber, B., Lehrach, H., and Adjaye, J. (2007). Fibroblast growth factor 2 modulates transforming growth factor beta signaling in mouse embryonic fibroblasts and human ESCs (hESCs) to support hESC self-renewal. Stem Cells 25, 455–464. Gurdon, J. B., and Melton, D. A. (2008). Nuclear reprogramming in cells. Science 322, 1811–1815. Hayashi, Y., Furue, M. K., Tanaka, S., Hirose, M., Wakisaka, N., Danno, H., Ohnuma, K., Oeda, S., Aihara, Y., Shiota, K., et al. (2010). BMP4 induction of trophoblast from mouse embryonic stem cells in defined culture conditions on laminin. In Vitro Cell. Dev. Biol. Anim. 46(5), 416–430. Heit, J. J., and Kim, S. K. (2004). Embryonic stem cells and islet replacement in diabetes mellitus. Pediatr. Diab. 5(Suppl. 2), 5–15. Hondo, E., and Stewart, C. L. (2005). Profiling gene expression in growth-arrested mouse embryos in diapause. Genome Biol. 6, 202. Humphrey, R. K., Beattie, G. M., Lopez, A. D., Bucay, N., King, C. C., Firpo, M. T., Rose-John, S., and Hayek, A. (2004). Maintenance of pluripotency in human embryonic stem cells is STAT3 independent. Stem Cells 22, 522–530. James, D., Levine, A. J., Besser, D., and Hemmati-Brivanlou, A. (2005). TGFbeta/activin/ nodal signaling is necessary for the maintenance of pluripotency in human embryonic stem cells. Development 132, 1273–1282. Janssens, S. (2007). Human embryonic stem cells for cardiac repair: The focus is on refined selection and cardiopoietic programming. Heart 93, 1173–1174. Jones, C. M., Kuehn, M. R., Hogan, B. L., Smith, J. C., and Wright, C. V. (1995). Nodalrelated signals induce axial mesoderm and dorsalize mesoderm during gastrulation. Development 121, 3651–3662. Kleinsmith, L. J., and Pierce, G. B., Jr. (1964). Multipotentiality of single embryonal carcinoma cells. Cancer Res. 24, 1544–1551. Kretzschmar, M., and Massague, J. (1998). SMADs: Mediators and regulators of TGF-beta signaling. Curr. Opin. Genet. Dev. 8, 103–111. Lawson, K. A., Meneses, J. J., and Pedersen, R. A. (1991). Clonal analysis of epiblast fate during germ layer formation in the mouse embryo. Development 113, 891–911. Lawson, K. A., Dunn, N. R., Roelen, B. A., Zeinstra, L. M., Davis, A. M., Wright, C. V., Korving, J. P., and Hogan, B. L. (1999). Bmp4 is required for the generation of primordial germ cells in the mouse embryo. Genes Dev. 13, 424–436. Levenstein, M. E., Ludwig, T. E., Xu, R. H., Llanas, R. A., VanDenHeuvel-Kramer, K., Manning, D., and Thomson, J. A. (2006). Basic fibroblast growth factor support of human embryonic stem cell self-renewal. Stem Cells 24, 568–574. Levine, A. J., and Brivanlou, A. H. (2006). GDF3, a BMP inhibitor, regulates cell fate in stem cells and early embryos. Development 133, 209–216. Levine, A. J., Levine, Z. J., and Brivanlou, A. H. (2009). GDF3 is a BMP inhibitor that can activate Nodal signaling only at very high doses. Dev. Biol. 325, 43–48. Ling, N., Ying, S. Y., Ueno, N., Shimasaki, S., Esch, F., Hotta, M., and Guillemin, R. (1986). Pituitary FSH is released by a heterodimer of the beta-subunits from the two forms of inhibin. Nature 321, 779–782. Lowry, W. E., Richter, L., Yachechko, R., Pyle, A. D., Tchieu, J., Sridharan, R., Clark, A. T., and Plath, K. (2008). Generation of human induced pluripotent stem cells from dermal fibroblasts. Proc. Natl. Acad. Sci. USA 105, 2883–2888.
Activin/Nodal Signaling and Pluripotency
55
Ludwig, T. E., Bergendahl, V., Levenstein, M. E., Yu, J., Probasco, M. D., and Thomson, J. A. (2006a). Feeder-independent culture of human embryonic stem cells. Nat. Methods 3, 637–646. Ludwig, T. E., Levenstein, M. E., Jones, J. M., Berggren, W. T., Mitchen, E. R., Frane, J. L., Crandall, L. J., Daigh, C. A., Conard, K. R., Piekarczyk, M. S., et al. (2006b). Derivation of human embryonic stem cells in defined conditions. Nat. Biotechnol. 24, 185–187. Martin, G. R. (1981). Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc. Natl. Acad. Sci. USA 78, 7634–7638. Matzuk, M. M., Kumar, T. R., Vassalli, A., Bickenbach, J. R., Roop, D. R., Jaenisch, R., and Bradley, A. (1995). Functional analysis of activins during mammalian development. Nature 374, 354–356. McLean, A. B., D’Amour, K. A., Jones, K. L., Krishnamoorthy, M., Kulik, M. J., Reynolds, D. M., Sheppard, A. M., Liu, H., Xu, Y., Baetge, E. E., et al. (2007). Activin a efficiently specifies definitive endoderm from human embryonic stem cells only when phosphatidylinositol 3-kinase signaling is suppressed. Stem Cells 25, 29–38. Mesnard, D., Guzman-Ayala, M., and Constam, D. B. (2006). Nodal specifies embryonic visceral endoderm and sustains pluripotent cells in the epiblast before overt axial patterning. Development 133, 2497–2505. Mishina, Y., Suzuki, A., Ueno, N., and Behringer, R. R. (1995). Bmpr encodes a type I bone morphogenetic protein receptor that is essential for gastrulation during mouse embryogenesis. Genes Dev. 9, 3027–3037. Miura, T., Luo, Y., Khrebtukova, I., Brandenberger, R., Zhou, D., Thies, R. S., Vasicek, T., Young, H., Lebkowski, J., Carpenter, M. K., et al. (2004). Monitoring early differentiation events in human embryonic stem cells by massively parallel signature sequencing and expressed sequence tag scan. Stem Cells Dev. 13, 694–715. Miyazono, K., Kusanagi, K., and Inoue, H. (2001). Divergence and convergence of TGFbeta/BMP signaling. J. Cell. Physiol. 187, 265–276. Moustafa, L. A., and Brinster, R. L. (1972). Induced chimaerism by transplanting embryonic cells into mouse blastocysts. J. Exp. Zool. 181, 193–201. Nakayama, T., Gardner, H., Berg, L. K., and Christian, J. L. (1998). Smad6 functions as an intracellular antagonist of some TGF-beta family members during Xenopus embryogenesis. Genes Cells 3, 387–394. Newman, M. B., and Bakay, R. A. (2008). Therapeutic potentials of human embryonic stem cells in Parkinson’s disease. Neurotherapeutics 5, 237–251. Nichols, J., Evans, E. P., and Smith, A. G. (1990). Establishment of germ-line-competent embryonic stem (ES) cells using differentiation inhibiting activity. Development 110, 1341–1348. Nichols, J., Chambers, I., Taga, T., and Smith, A. (2001). Physiological rationale for responsiveness of mouse embryonic stem cells to gp130 cytokines. Development 128, 2333–2339. Niwa, H., Burdon, T., Chambers, I., and Smith, A. (1998). Self-renewal of pluripotent embryonic stem cells is mediated via activation of STAT3. Genes Dev. 12, 2048–2060. Okita, K., Hong, H., Takahashi, K., and Yamanaka, S. (2010). Generation of mouseinduced pluripotent stem cells with plasmid vectors. Nat. Protoc. 5, 418–428. Ou, L., Wang, X., and Zou, F. (2010). Is iPS cell the panacea? IUBMB Life 62, 170–175. Park, I. H., Arora, N., Huo, H., Maherali, N., Ahfeldt, T., Shimamura, A., Lensch, M. W., Cowan, C., Hochedlinger, K., and Daley, G. Q. (2008a). Disease-specific induced pluripotent stem cells. Cell 134, 877–886. Park, I. H., Lerou, P. H., Zhao, R., Huo, H., and Daley, G. Q. (2008b). Generation of human-induced pluripotent stem cells. Nat. Protoc. 3, 1180–1186.
56
Zhenzhi Chng et al.
Rebagliati, M. R., Toyama, R., Fricke, C., Haffter, P., and Dawid, I. B. (1998). Zebrafish nodal-related genes are implicated in axial patterning and establishing left-right asymmetry. Dev. Biol. 199, 261–272. Reynolds, D., Vallier, L., Chng, Z., and Pedersen, R. A. (2009). Signalling pathways in embryonic stem cells. In “Regulatory Networks in Stem Cells,” (V. K. Rajasekhar and M. C. Vemuri, Eds.). Humana Press, New York, NY, pp. 293–308. Robertson, E. J., Norris, D. P., Brennan, J., and Bikoff, E. K. (2003). Control of early anterior-posterior patterning in the mouse embryo by TGF-beta signalling. Philos. Trans. R. Soc. Lond. B Biol. Sci. 358, 1351–1357, discussion 1357. Ronaghi, M., Erceg, S., Moreno-Manzano, V., and Stojkovic, M. (2010). Challenges of stem cell therapy for spinal cord injury: Human embryonic stem cells, endogenous neural stem cells, or induced pluripotent stem cells? Stem Cells 28, 93–99. Rosler, E. S., Fisk, G. J., Ares, X., Irving, J., Miura, T., Rao, M. S., and Carpenter, M. K. (2004). Long-term culture of human embryonic stem cells in feeder-free conditions. Dev. Dyn. 229, 259–274. Schier, A. F. (2003). Nodal signaling in vertebrate development. Annu. Rev. Cell Dev. Biol. 19, 589–621. Smith, A. G., and Hooper, M. L. (1987). Buffalo rat liver cells produce a diffusible activity which inhibits the differentiation of murine embryonal carcinoma and embryonic stem cells. Dev. Biol. 121, 1–9. Smith, A. G., Heath, J. K., Donaldson, D. D., Wong, G. G., Moreau, J., Stahl, M., and Rogers, D. (1988). Inhibition of pluripotential embryonic stem cell differentiation by purified polypeptides. Nature 336, 688–690. Smith, J. C., Price, B. M., Van Nimmen, K., and Huylebroeck, D. (1990). Identification of a potent Xenopus mesoderm-inducing factor as a homologue of activin A. Nature 345, 729–731. Smith, J. R., Vallier, L., Lupo, G., Alexander, M., Harris, W. A., and Pedersen, R. A. (2008). Inhibition of Activin/Nodal signaling promotes specification of human embryonic stem cells into neuroectoderm. Dev. Biol. 313, 107–117. Solter, D., Skreb, N., and Damjanov, I. (1970). Extrauterine growth of mouse egg-cylinders results in malignant teratoma. Nature 227, 503–504. Stevens, L. C. (1970). The development of transplantable teratocarcinomas from intratesticular grafts of pre- and postimplantation mouse embryos. Dev. Biol. 21, 364–382. Stevens, L. C., and Little, C. C. (1954). Spontaneous testicular teratomas in an inbred strain of mice. Proc. Natl. Acad. Sci. USA 40, 1080–1087. Takahashi, K., and Yamanaka, S. (2006). Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126, 663–676. Takahashi, K., Tanabe, K., Ohnuki, M., Narita, M., Ichisaka, T., Tomoda, K., and Yamanaka, S. (2007). Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131, 861–872. Tesar, P. J., Chenoweth, J. G., Brook, F. A., Davies, T. J., Evans, E. P., Mack, D. L., Gardner, R. L., and McKay, R. D. (2007). New cell lines from mouse epiblast share defining features with human embryonic stem cells. Nature 448, 196–199. Thomson, J. A., and Marshall, V. S. (1998). Primate embryonic stem cells. Curr. Top. Dev. Biol. 38, 133–165. Thomson, J. A., Itskovitz-Eldor, J., Shapiro, S. S., Waknitz, M. A., Swiergiel, J. J., Marshall, V. S., and Jones, J. M. (1998). Embryonic stem cell lines derived from human blastocysts. Science 282, 1145–1147. Valdimarsdottir, G., and Mummery, C. (2005). Functions of the TGFbeta superfamily in human embryonic stem cells. APMIS 113, 773–789.
Activin/Nodal Signaling and Pluripotency
57
Vale, W., Rivier, J., Vaughan, J., McClintock, R., Corrigan, A., Woo, W., Karr, D., and Spiess, J. (1986). Purification and characterization of an FSH releasing protein from porcine ovarian follicular fluid. Nature 321, 776–779. Vallier, L., and Pedersen, R. A. (2005). Human embryonic stem cells: An in vitro model to study mechanisms controlling pluripotency in early mammalian development. Stem Cell Rev. 1, 119–130. Vallier, L., Reynolds, D., and Pedersen, R. A. (2004). Nodal inhibits differentiation of human embryonic stem cells along the neuroectodermal default pathway. Dev. Biol. 275, 403–421. Vallier, L., Alexander, M., and Pedersen, R. A. (2005). Activin/Nodal and FGF pathways cooperate to maintain pluripotency of human embryonic stem cells. J. Cell Sci. 118, 4495–4509. Vallier, L., Mendjan, S., Brown, S., Chng, Z., Teo, A., Smithers, L. E., Trotter, M. W., Cho, C. H., Martinez, A., Rugg-Gunn, P., et al. (2009a). Activin/Nodal signalling maintains pluripotency by controlling Nanog expression. Development 136, 1339–1349. Vallier, L., Touboul, T., Brown, S., Cho, C., Bilican, B., Alexander, M., Cedervall, J., Chandran, S., Ahrlund-Richter, L., Weber, A., et al. (2009b). Signaling pathways controlling pluripotency and early cell fate decisions of human induced pluripotent stem cells. Stem Cells 27, 2655–2666. Vallier, L., Touboul, T., Chng, Z., Brimpari, M., Hannan, N., Millan, E., Smithers, L. E., Trotter, M., Rugg-Gunn, P., Weber, A., et al. (2009c). Early cell fate decisions of human embryonic stem cells and mouse epiblast stem cells are controlled by the same signalling pathways. PLoS ONE 4, e6082. Varlet, I., Collignon, J., and Robertson, E. J. (1997). Nodal expression in the primitive endoderm is required for specification of the anterior axis during mouse gastrulation. Development 124, 1033–1044. Verlinsky, Y., Strelchenko, N., Kukharenko, V., Rechitsky, S., Verlinsky, O., Galat, V., and Kuliev, A. (2005). Human embryonic stem cell lines with genetic disorders. Reprod. Biomed. Online 10, 105–110. Williams, R. L., Hilton, D. J., Pease, S., Willson, T. A., Stewart, C. L., Gearing, D. P., Wagner, E. F., Metcalf, D., Nicola, N. A., and Gough, N. M. (1988). Myeloid leukaemia inhibitory factor maintains the developmental potential of embryonic stem cells. Nature 336, 684–687. Winnier, G., Blessing, M., Labosky, P. A., and Hogan, B. L. (1995). Bone morphogenetic protein-4 is required for mesoderm formation and patterning in the mouse. Genes Dev. 9, 2105–2116. Wrighton, K. H., Lin, X., Yu, P. B., and Feng, X. H. (2009). Transforming growth factor beta can stimulate Smad1 phosphorylation independently of bone morphogenic protein receptors. J. Biol. Chem. 284, 9755–9763. Xia, Y., and Schneyer, A. L. (2009). The biology of activin: Recent advances in structure, regulation and function. J. Endocrinol. 202, 1–12. Xiao, L., Yuan, X., and Sharkis, S. J. (2006). Activin A maintains self-renewal and regulates fibroblast growth factor, Wnt, and bone morphogenic protein pathways in human embryonic stem cells. Stem Cells 24, 1476–1486. Xu, R. H. (2006). In vitro induction of trophoblast from human embryonic stem cells. Methods Mol. Med. 121, 189–202. Xu, R. H., Sampsell-Barron, T. L., Gu, F., Root, S., Peck, R. M., Pan, G., Yu, J., Antosiewicz-Bourget, J., Tian, S., Stewart, R., et al. (2008). NANOG is a direct target of TGFbeta/activin-mediated SMAD signaling in human ESCs. Cell Stem Cell 3, 196–206.
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Yamamoto, M., Saijoh, Y., Perea-Gomez, A., Shawlot, W., Behringer, R. R., Ang, S. L., Hamada, H., and Meno, C. (2004). Nodal antagonists regulate formation of the anteroposterior axis of the mouse embryo. Nature 428, 387–392. Yao, S., Chen, S., Clark, J., Hao, E., Beattie, G. M., Hayek, A., and Ding, S. (2006). Longterm self-renewal and directed differentiation of human embryonic stem cells in chemically defined conditions. Proc. Natl. Acad. Sci. USA 103, 6907–6912. Ying, Q. L., Nichols, J., Chambers, I., and Smith, A. (2003). BMP induction of Id proteins suppresses differentiation and sustains embryonic stem cell self-renewal in collaboration with STAT3. Cell 115, 281–292. Yu, J., Vodyanik, M. A., Smuga-Otto, K., Antosiewicz-Bourget, J., Frane, J. L., Tian, S., Nie, J., Jonsdottir, G. A., Ruotti, V., Stewart, R., et al. (2007). Induced pluripotent stem cell lines derived from human somatic cells. Science 318, 1917–1920. Zhou, H., Wu, S., Joo, J. Y., Zhu, S., Han, D. W., Lin, T., Trauger, S., Bien, G., Yao, S., Zhu, Y., et al. (2009). Generation of induced pluripotent stem cells using recombinant proteins. Cell Stem Cell 4, 381–384.
C H A P T E R
F O U R
Intracrine Signaling Mechanisms of Activin A and TGF-b Olav A. Gressner Contents I. Common TGF-b and Activin A Signaling and Target Genes: Focusing on CTGF/CCN2 II. The Early Response to Cellular Stress: Intracellular Activation of TGF-b III. Continuous Low-Level Activation of Activin A/TGF-b Target Genes IV. Intracrine Signaling: General Aspects V. Intracrine Signaling of TGF-b: The Stimulatory Pathway VI. Intracrine Signaling of TGF-b: The Inhibitory Pathway VII. Intracrine Activin A Signaling VIII. Conclusion References
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Abstract While autocrine stimulation of cells by Activin A and/or its family member transforming growth factor b (TGF-b) is a phenomenon observed in a variety of cell types, little is known of putative intracellular signaling loops of these cytokines. Intracellular actions of several peptide hormones, growth factors, as well as of extracellular signaling enzymes and DNA-binding proteins, either within target cells or within their cells of synthesis have been shown. Although these intracrine moieties are structurally diverse, they share certain characteristics of synthesis and function. Depending on the cell type, there are reports of stimulatory as well as inhibitory mechanisms induced by such intracrine mechanisms, and this also accounts for transforming growth factor b (TGF-b), whereas only stimulatory intracrine signaling of Activin A could be demonstrated so far. Stimulatory intracrine signaling loops of TGF-b were shown following calpain-dependent intracellular proteolytic activation of the latent cytokine in hepatocytes under cellular stress conditions of this cytokine, leading to transcriptional activation of connective tissue growth factor (CTGF/CCN2)
Wisplinghoff Medical Laboratories, Classen-Kappelmann Str. 24, Cologne, Germany Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00004-4
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2011 Elsevier Inc. All rights reserved.
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as a representative TGF-b-sensitive reporter gene. In contrast to TGF-b, increasing intrahepatocellular concentrations of Activin A are not the result of release from an intracellularly deposited latent complex, but of active de novo synthesis. The stimulatory intracrine signaling pathways of both, TGF-b and Activin A, are proposed to occur via Alk4/Alk5 receptors and Smad2, whereas additional activation of Smad3 only seems to be involved in intracrine Activin A signaling. However, intracrine TGF-b signaling may itself also be inhibitory as active TGF-b is also able to bind to intracellular TGF-b type II receptor, resulting in a ligandinduced impediment in receptor trafficking to the cell surface. Whether stimulatory or inhibitory modulation of the TGF-b pathway takes place seems to depend on the cell type and environmental conditions. Future studies are necessary at this point. ß 2011 Elsevier Inc.
I. Common TGF-b and Activin A Signaling and Target Genes: Focusing on CTGF/CCN2 The transforming growth factor b (TGF-b) superfamily of growth factors includes more than 30 structurally related proteins with diverse functions in embryonic development and adult tissue homeostasis. They can be grouped into three major families: the TGF-b family, the Activin family, and the bone morphogenetic protein (BMP) family. TGF-b was the first member of the superfamily to be isolated, and was initially recognized by its capacity to induce anchorage-independent proliferation of rat fibroblasts (Roberts et al., 1981). Such as Activin, TGF-b itself acts by binding to specific transmembraneous serine/threonine kinase receptors, which induce intracellular signaling by phosphorylation of receptor-regulated Smads, so-called R-Smads, which after nuclear translocation initiate target gene transcription (ten Dijke et al., 2000). Although Activin binds to its own specific receptors (Alk4), the same set of Smads that is used by TGF-b itself (Smad2/3) is recruited for its signal transduction. This TGF-b/Activin signaling is attenuated by Smad7, an inhibitory Smad (I-Smad) (Miyazawa et al., 2002; Pangas and Woodruff, 2000). As TGF-b and Activins generate signaling by common intracellular mediators, their set of activated target genes is in many cases overlapping (Roberts et al., 1991). One common TGF-b and Activin A target gene is connective tissue growth factor (CTGF/CCN2). CTGF is a cysteine-rich, matrix-associated, heparin-binding protein which in vitro mirrors some of the effects of TGF-b on skin fibroblasts, such as stimulation of extracellular matrix production, proliferation, chemotaxis, and integrin expression. By doing so, CTGF was shown to promote endothelial cell growth, migration, adhesion, and survival and is thus implicated in endothelial cell function and angiogenesis (Moussad and Brigstock, 2000). As CTGF is furthermore capable of upregulating both matrix metalloproteinases (MMPs) and their inhibitors
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(TIMPs), involvement in extracellular matrix remodeling in wound healing, scleroderma, and other fibrotic processes has been suggested (Moussad and Brigstock, 2000). Even though the expression of CTGF is equally controlled by both Activin A and TGF-b, the role of Smad2 and Smad3 in the regulation of the CTGF promoter seems to be largely cell type specific (Gressner and Gressner, 2008; Grotendorst, 1997; Leask and Abraham, 2006; Leask et al., 2003; Moussad and Brigstock, 2000). For example, Smad2 seems to be the major inductor of CTGF gene transcription in hepatocytes (Gressner et al., 2009), while dermal fibroblasts demonstrate a Smad3-dependent regulation of the CTGF promoter (Leask et al., 2001, 2003).
II. The Early Response to Cellular Stress: Intracellular Activation of TGF-b TGF-b is a 25-kDa homodimeric cytokine derived by intracellular proteolytic processing of a larger proprotein (Massague, 1998). However, once cleaved from the cytokine, a dimer of the glycosylated N-terminal prosegment of TGF-b, called the latency-associated peptide (LAP), remains noncovalently associated with the mature TGF-b after secretion (Koli et al., 2001). This complex of TGF-b and LAP, the small latent complex (SLC), is functionally inactive, and the dissociation of TGF-b from LAP is a central regulatory step in the modulation of TGF-b actions. The SLC can regroup as a large latent complex (LLC) through bondage of cysteines in the LAP with a pair of cysteines in the third 8-cys domain of the latent TGF-b binding protein (LTBP) (Gleizes et al., 1996; Saharinen et al., 1996). TGF-b itself is secreted by most cell types as part of this LLC in a biologically inactive form (Koli et al., 2001). Once extracellular, active (mature) TGF-b is then released from the LLC by several proteins, including the multifunctional glycoprotein thrombospondin-1, calpains, plasmin, and some integrins (Koli et al., 2001). Thus, conversion from the latent precursor molecule to the biologically active, mature form, and initiation of the phosphorylation of the intracellular cascade of Smad proteins (Inagaki and Okazaki, 2007) is a key step in the regulation of TGF-b biological activity in vivo and in vitro (Fig. 4.1). Of note, the presence of several components of the TGF-b system such as LAP (Roth-Eichhorn et al., 1998), LTBP (Roth et al., 1997), and TGF-b itself was also shown in the cytoplasm of cultured hepatocytes and other cells (Roth et al., 1998; Roth-Eichhorn et al., 1998). The colocalization of TGF-b, LAP, and LTBP in the microsomal fraction, also containing endosomal and Golgi vesicles, suggests an uptake and metabolism of latent TGF-b by hepatocytes. It is well established on mRNA level that normal
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Mannose-6- P -residues
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Figure 4.1 Schematic model of synthesis, secretion, and matrix deposition of TGF-b. TGF-b is synthesized as a preproprotein, which then is proteolytically processed in the rough endoplasmic reticulum. Two monomers of TGF-b dimerize through disulfide bridges, and the pro-TGF-b dimer is then cleaved by the furin convertase to yield the small latent TGF-b complex (SLC), in which the latency-associated peptide (LAP) and the mature peptide are connected by noncovalent bonds. The large latent TGF-b complex (LLC) is formed by covalent attachment of the large latent TGF-b binding protein (LTBP). The LLC of TGF-b is secreted by exocytosis to the extracellular region. The N-terminal and hinge region of LTBP interact with components of the extracellular matrix which can be covalent owing to crosslinking by transglutaminase. As TGF-b in the LLC is functionally inactive and cannot interact with the receptors, it requires activation, meaning release from the LLC. This may be achieved by many mechanisms: thrombospondin-1 binding to the LLC, integrin alphaV-b6 binding to the LLC, proteolytic cleavage of the LTBPs, reactive oxygen species, or low pH. The release of mature dimeric TGF-b is essentially a mechanical process that demands cleavage and opening of the LLC structure so that the caged mature C-terminal TGF-b polypeptide is released to reach the receptor. Mature TGF-b can then bind to its cognate receptors, type III, type II, and Alk5 TGF-b type I receptor, leading to intracellular Smad2/3 activation.
hepatocytes do not express de novo any of the TGF-b isoforms (Dallas et al., 1994; De Bleser et al., 1997). However, the localization of TGF-b and LAP but not LTBP in central cellular parts such as perinuclear and in mitochondria, and the enrichment of LTBP near the hepatocytes plasma membrane proposed also a separate existence of the proteins, which might be a result of intracellular cleavage of the LLC, possibly leading to intracellular TGF-b activation. The increasingly intense staining for active (mature) TGF-b of those hepatocytes that have lost the majority of cell–cell contacts suggests the
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importance of intact cell formations or intercellular signals in the prevention of intracellular TGF-b activation. In summary, as hepatocytes seem to be able of internalize and deposit the LLC, they have been put in the focus of research on a possible intracellular activation and signaling of TGF-b. Based on previous data from this laboratory (Gao et al., 1996; Gressner and Wulbrand, 1997), recent investigations have focused on calpains as intracellularly acting proteases potentially responsible for activation of latent TGF-b in hepatocytes. Calpains consist of many types of calcium-dependent cytoplasmic proteases, which differ in their requirement for free Ca2þ (Mehendale and Limaye, 2005). Predominantly, they are in an inactive form, but several cellular events can increase intracellular calpain activity such as rise of cytosolic free Ca2þ, dissociation from the membrane, and decreases in the level of calpastatin, their endogenous inhibitor (Mehendale and Limaye, 2005). Current knowledge suggests that some of these events take place in isolated and cultured hepatocytes and in injured livers as response to cellular stress (Arora et al., 1996; Limaye et al., 2003; Fig. 4.2). This assumption is supported by our earlier findings that inactivation of calpains attenuates spontaneous apoptosis of hepatocytes (Gressner et al., 1997). Furthermore, calpains were shown to be involved in integrinmediated signal transduction (Inomata et al., 1996) which is greatly disturbed during hepatocyte isolation in vitro and in the injured liver in vivo. In confirmation of this, we have previously shown that the composition of culture matrix determines the staining intensity of intracellular TGF-b in hepatocytes, which correlates inversely with the hepatocellular phenotype (Gressner and Wulbrand, 1997), suggesting integrin-linked signals as modulators of intracellular TGF-b activation and of survival of hepatocytes (Gkretsi et al., 2007). Even the release of calpains from dying hepatocytes is suggested as a mechanism, which mediates progression of acute liver injury (Limaye et al., 2003). So, taken together, several reports point to a significant role of calpain activation in the early stage of liver injury and acute hepatocellular stress (Arora et al., 1996).
III. Continuous Low-Level Activation of Activin A/TGF-b Target Genes As discussed above, CTGF expression in hepatocytes, but not only in this cell type, is efficiently stimulated by exogenous TGF-b. Also, Activin A was identified as an inducer of CTGF synthesis in hepatocytes (Gressner et al., 2008a). Signal transduction of exogenous TGF-b and/or Activin A to the CTGF promoter follows the common pathway of intracellular signaling, in which phosphorylation of Activin receptor-like kinases (Alk4/5) and
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Mouse IgG (0 h)
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Figure 4.2 Time course of APAAP immunostaining of TGF-b in cultured hepatocytes. (A) APAAP immunostaining of TGF-b in freshly isolated hepatocytes (0 h) and in hepatocytes cultured for 24 h under serum-free conditions. A monoclonal antibody against the three isoforms of active (not latent) TGF-b was used. Controls were performed with nonspecific mouse immunoglobulin G (IgG) instead of TGF-b specific first antibody (original magnification 40). (B) APAAP immunostaining of TGF-b in hepatocytes cultured for 24 h under serum-free conditions but in the presence or absence of cycloheximide (5 mM) (original magnification 40).
Smad2/3 plays an important role for CTGF expression (Gressner et al., 2007, 2008a,b, 2009; Miyazawa et al., 2002; Pangas and Woodruff, 2000). However, several studies have pointed to a continuous basal (low level) transcriptional activation of TGF-b/Activin A target genes (Gressner et al., 2008a,b; Jachimczak et al., 1996). The CAGA-luc reporter gene, a TGF-b target sequence in the DNAbinding protein inhibitor (ID-1) promoter, displays low basal activity when transfected into hepatocytes (Eijken et al., 2007), even when these cells are cultured in complete absence of exogenous TGF-b and/or Activin A. Similar accounts for the CTGF promoter, whose basal activity may furthermore be reduced in the presence of intracellularly acting calpain inhibitors (Gressner et al., 2008b). These findings propose putative intracellular loops for TGF-b and/or Activin A in hepatocytes, leading to a low-level target gene activation, and it may be hypothesized that intrinsic activation of latent TGF-b in hepatocytes (and other cell types), and subsequent signaling to TGF-b-responsive target genes could be an immediate or early response of the cell to cultural and injurious stress (Fig. 4.3).
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Figure 4.3 Effect of external and internal inhibitors of TGF-b signaling on CTGF expression in hepatocytes cultured under TGF-b-free conditions. (A) Western blot of CTGF of hepatocytes cultured for 24 h under completely serum-free conditions with or without addition of 5 ng/ml rhTGF-b1. Some cultures received the Alk4/5 inhibitor (SB-431542, 5 mM), lysates were probed for CTGF/CCN2 and b-actin expression. A representative blot of four independent experiments is shown. (B) Immunoprecipitation of metabolically labeled CTGF. Hepatocytes were cultured in serum-, cysteineand methionine-free DMEM for 24 h in the absence or presence of Alk4/5 inhibitor (SB-431542, 5 mM) and labeled with the PRO-MIX L-[35S] methionine/cysteine in vitro cell labeling kit each for 3 h at indicated time points. (C) Western blot of CTGF of hepatocytes cultured for 24 h under complete serum-free conditions. Cultures received various inhibitors of extra- and intracellular TGF-b1 signaling like rsTbRII (1 mg/ml), neutralizing anti-TGF-b1 antibody (4 mg/ml) and rhLAP [TGF-b] (1 mg/ ml). Lysates were probed for CTGF/CCN2 and b-actin expression. A representative blot of three independent experiments is shown. (D) CTGF/CCN2 reporter gene activation. Hepatocytes were prepared as described in (B), but received a blocking anti-hTbRII antibody (10 mg/ml). They were cultured for 24 h under complete serumfree conditions. CTGF-luciferase activities are shown relative to Renilla luciferase activity. Mean values SD of four experiments are shown.
IV. Intracrine Signaling: General Aspects It has only been a couple of years that theories of intracellular hormone/cytokine action have emerged which was designated as “intracrine” signaling. Intracrine signaling describes the action of an actually extracellularly acting signaling peptide either after internalization by the respective cell or by retention in the cell after synthesis (Re and Cook, 2006). Intracellular actions of several (peptide) hormones, growth factors, DNA-binding proteins, or extracellular signaling enzymes, either within the cell of their synthesis or within specific target cells, have been shown (Re and
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Cook, 2006). Although these intracrine moieties are structurally diverse, they share certain characteristics of synthesis and function. In addition to this, emerging data are strongly supportive of the notion that intracellular intracrine actions are physiologically relevant, and, thus, may be considered as a target for pharmacologic intervention (Re and Cook, 2006, 2007, 2008).
V. Intracrine Signaling of TGF-b: The Stimulatory Pathway We discussed above that latent TGF-b may be activated intracellularly and that it is able to trigger a signaling cascade within the hepatocyte leading to target gene (e.g., CTGF) expression. However, the type of signaling pathway activated by intracellularly activated TGF-b was not yet defined. Two hypotheses can be suggested: (i) the intracellularly deposited LLC is released from the cell and TGF-b is activated extracellularly, then binding to membrane-bound TGF-b type I and type II receptors and thus, inducing the “classical” Smad signaling pathway that leads to target gene expression (autocrine stimulation). (ii) TGF-b is released from the LLC within the cell and triggers entirely intracellular pathways leading to target gene expression (intracrine stimulation). Both mechanisms have been proposed in the literature (Fernandez et al., 2002; Gressner et al., 2008b; Jachimczak et al., 1996; Koli et al., 2001), and it is very likely that both of them occur, depending on the cell type or the environmental conditions investigated. By performing antisense experiments, Jachimczak et al. could show that glioma cell proliferation is stimulated in part by endogenously produced TGF-b via an intracrine loop mechanism. Similar to that observed by our laboratory, extracellularly acting, neutralizing TGF-b antibodies exerted only moderate effects in approximately 40% of all cells. The remaining 60% were therefore likely stimulated in an autocrine external loop mechanism ( Jachimczak et al., 1996). Results from our laboratory that were obtained in hepatocytes, which were cultured in conditions entirely free of exogenous TGF-b, showed that following intrahepatocellular activation of latent TGF-b by calpains, Smad2 but not Smad3 phosphorylation was induced, using an entirely intracrine interaction of the cytokine with the Alk5 receptor/TGF-b type I receptor (Gressner et al., 2008b). However, in contrast to what was described for glioma cells, the possibility of an autocrine extracellular stimulation of the cells by previously released latent and then activated TGF-b was basically entirely excluded in hepatocytes, as a variety of noncell-permeable inhibitors of extracellular TGF-b signaling, such as soluble TGF-b type II receptor or neutralizing human TGF-b type II receptor antibody, as well as extracellular TGF-b scavengers like human LAP [TGF-b] or neutralizing anti-TGF-b antibody, did not have a diminishing effect on TGF-b target gene
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(i.e., CTGF) expression at all (Gressner et al., 2008b). Only the application of an inhibitor of the intracellular kinase domain of the Alk4/5 receptor (SB431542) (Inman et al., 2002) resulted in a significant reduction of basal CTGF expression, triggered by intracellular TGF-b (Gressner et al., 2008b). Thus, as stimulatory intracellular TGF-b signaling does not seem to involve ligand binding to the cell-surface TGF-b type II receptor, which is in contrast to extracellular TGF-b signaling, but seems to be premised on a direct interaction of the endogenously activated cytokine with the intracellular domain of its type I receptor, two possible mechanisms of interaction between these two partners (TGF-b/TGF-b type I receptor) may be discussed: (i) intracellularly activated TGF-b binds to its original binding site on the extracellular domain of the yet intracellular, soluble Alk5/TGF-b type I receptor and, thereby, activates its serine–threonine kinase. Or (ii) TGF-b binds to a not yet identified binding site at the intracellular domain of the membranous Alk5 receptor and, by doing so, causes kinase activation through accessory pathways. Even though the latter hypothesis seems unlikely as ligand-bound receptor complexes are considered unable to recruit cell surface adaptor proteins such as the Smad anchor for receptor activation (SARA), which in turn are required for the phosphorylation of Smad2 (Tsukazaki et al., 1998), further investigations are certainly necessary at this point (Fig. 4.4). Trafficking of ligands and receptors is nowadays considered essential in regulating both intensity and time course of signaling transduction. In a steady state, cells produce receptors continuously and, to maintain homeostasis, also need to degrade the previously membrane-bound receptors at a certain rate. Degradation usually takes place in intracellular vesicular compartments such as lysosomes, but receptors can escape this fate by trafficking back to the plasma membrane (recycling). Such a complex intracellular trafficking raises the possibility that signal transduction events, originally downstream of the receptor, do not necessarily have to take place at the plasma membrane but also en route, while the ligand/receptor complex traffics through specific intracellular, endosomal compartments (GonzalezGaitan, 2008). This would furthermore strengthen hypothesis (i). Of note, TGF-b receptors are constitutively internalized (Tan et al., 2004; Wiley et al., 2003), but the trafficking route that the receptors follow thereafter is highly depending on whether they are part of a signaling complex or not. And in turn, different routes will trigger different signaling pathways and furthermore affect how or if receptors are subject to degradation. It is interesting that all data that are currently existing point to a neglectable role of TAK1- (and MAP-kinase) activation in the process of stimulatory intracellular TGF-b signaling. Even to the contrary, significantly enhanced hepatocellular CTGF expression was observed following specific inhibition of TAK1 by 5-Z-7-oxozeaenol (unpublished data by the author), which could be the result of a stimulation of the cell by other cytokines. TAK1 is found attached to the intracellular domain not only of
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Figure 4.4 Two hypotheses of intracellular Alk5 stimulation during stimulatory intracellular TGF-b signaling. Two possible mechanisms of interaction between these two partners may be discussed: (i) intracellularly activated TGF-b binds to its original binding site on the extracellular domain of the yet intracellular, soluble Alk5/TGF-b type I receptor and, thereby, activates its serine–threonine kinase. Or (ii) TGF-b binds to a not yet identified binding site at the intracellular domain of the membranous Alk5 receptor and causes kinase activation through accessory pathways.
the TGF-b type II receptor but also of Alk3, the constitutive receptor for BMP2/4/7 signaling (Chen et al., 2004; Nohe et al., 2004) and is activated upon binding of BMP4 (Nohe et al., 2004), suggesting that the increase in spontaneous CTGF expression following TAK1 inhibition could be the result of an autocrine stimulation of the Alk3-receptor by BMP2/4/7 (Fig. 4.5). Expanding on this idea, the finding of TAK1-mediated inhibition of intracrine TGF-b signaling to the CTGF promoter could indicate a BMP2/4/7-dependent and Alk3-receptor-mediated antagonism against
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Figure 4.5 A simplified and schematic overview of the proposed activation process of latent TGF-b within hepatocytes and of its intracellular signaling triggering TGF-b target gene (e.g., CTGF) gene expression under consideration of the current literature. Latent TGF-b, which is deposited within the hepatocyte, is activated by proteases such as calpain within the first 24 h following culture-induced stress or in vivo injury of the liver parenchyma. Via intracellular activation of the kinase domain of the Alk5 receptor, it triggers Smad2phosphorylation but not Smad3-phosphorylation leading to nuclear translocation of the first Smad together with the common mediator Smad4 and subsequent TGF-b target gene (e.g., CTGF) expression. Even though still hypothetical, there are hints of a BMP2/4/7-dependent and Alk3-receptor-mediated antagonism against intracellular TGF-b signaling during early liver impairment, possibly via TAK1-mediated transcriptional upregulation of Smad7. TAK1 is found attached to the intracellular domain not only of the TGF-b type II receptor but also of Alk3, the constitutive receptor for BMP2/4/7 signaling, and is activated upon binding of BMP4. SARA, Smad anchor for receptor activation.
intracellular TGF-b signaling during early liver impairment, possibly via TAK1-mediated transcriptional upregulation of inhibitory Smad7 (Dowdy et al., 2003; Whitman, 1997). Supportive of that, hepatocellular Smad7 was found upregulated during acute liver injury (Seyhan et al., 2006) and also intrahepatic BMP4 was reported to be upregulated in the bile duct ligation model of the rat, here resulting in detectable Smad1 phosphorylation (Fan et al., 2006). Of note, Smad1 is the classical target Smad of BMP2/4/7dependent Alk3 activation (Nohe et al., 2004).
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VI. Intracrine Signaling of TGF-b: The Inhibitory Pathway In the previous chapter, we talked about the ability of TGF-b to stimulate the Smad signaling pathway through intracrine action and interaction with the TGF-b type I (Alk4/5) receptor. However, there is also evidence that TGF-b is able to display inhibitory actions through intracrine signaling (Fig. 4.6). In their 2002 study in plasmocytoma cell lines, Fernandez et al. demonstrated that the generation of active TGF-b within a cell is able to disrupt autocrine TGF-b signaling via the formation of nonproductive intracellular ligand–receptor complexes. This confirms previous reports on a functionally relevant role of intracellular ligand–receptor interactions. For example, acquired defects in the membrane localization of growth factor receptors have previously been demonstrated in cells transformed by the viral oncoprotein v-sis (Aaronson et al., 1986; Fleming et al., 1989). In these studies, the diminution of platelet-derived growth factor receptor presentation at the cell surface was mechanistically linked to internal activation of the
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Figure 4.6 Extracellular activation and signaling of TGF-b and its autoregulatory inhibition through inhibitory intracellular signaling of the same cytokine. (A) Extracellular activation and signaling of TGF-b: Latent TGF-b is activated outside the cell where it can bind to the cell surface TGF-b type II receptor which, after dimerization with the TGF-b type I receptor, induces Smad2/3 phosphorylation. (B) Inhibitory intracellular signaling of TGF-b: In plasmocytoma cell lines, Fernandez et al. demonstrated that TGF-b is activated already within the cell and can as such bind the cytoplasmic (not yet membrane bound) TGF-b type II receptor, thereby preventing its trafficking to the cell surface. SARA, Smad anchor for receptor activation; TbRI, TGF-b type I receptor; TbRII, TGF-b type II receptor; TbRIII, TGF-b type III receptor; TGFBR2, TGF-b type II receptor gene.
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receptor by v-sis. However, although intracellular autocrine loops have been described for v-sis itself such as for other cytokines (Bejcek et al., 1989; Browder et al., 1989), the data published by Fernandez et al. suggest that intracellular ligand–receptor complexes of TGF-b are not necessarily capable of initiating signaling. Much more, they found that intracellular sequestration of the TGF-b type II receptor by activated or active TGF-b prevents the receptor from trafficking to the cell surface, thereby reducing cell surface presentation of the type II receptor, which is necessary for exogenously initiated TGF-b signaling (Fig. 4.7). The preponderance of either such inhibitory intracrine pathways or stimulatory intracrine pathways as described in the previous chapter may, at least partially, be due to cell type or environmental-specific differences in the trafficking behavior of TGF-b type I receptor and type II receptors
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Figure 4.7 Effect of various inhibitors of intra- and extracellular Activin A signaling as well as of knockdown of inhibin bA by RNA interference in hepatocytes cultured under Activin A-free conditions. Western blots of CTGF (A) as well as phosphorylated, and total Smads 1,2 and 3 (B) of hepatocytes cultured for 24 h in serum-free conditions. Cultures received the neutralizing a-Activin A antibody (2 mg/ml), follistatin (1 mg/ml), or the intracellularly acting Alk4/5-inhibitor (5 mM; SB431542). Lysates were probed for CTGF/ CCN2, p-Smads 1/2/3, total Smad2/3, and b-actin expression, respectively. A representative blot of three independent experiments is shown. (C) Hepatocytes were transfected with pSilencer small interfering RNAs (siRNAs) directed against the inhibin bA gene encoding the bA subunit of Activin A. (I) Efficiency of inhibin bA-knockdown was proven by RT-PCR (II). Protein lysates were probed for CTGF (III) or p-Smad2 expression by Western blot analysis. All experiments are representative for three independent transfections. Co, control; si, siRNA; mock, mock siRNA.
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(Wells et al., 1997). For example, Wu and Derynck only recently gave evidence that glucose overabundance leads to a mobilization of an intracellular pool of TGF-b type I and type II receptors, and that glucose itself acts as a stimulator of their trafficking to and their integration into the cell membrane. By this, glucose is able to rapidly induce cell surface presentation of both receptors without increasing their expression levels (Wu and Derynck, 2009). In conclusion, next to the known stimulatory actions of the intracellular ligand, these data demonstrate how the activation of TGF-b within a cell can impact the TGF-b receptor presence on the cell surface, and therefore also negatively impact the responsiveness to both autocrine and paracrine sources of the cytokine.
VII. Intracrine Activin A Signaling We previously discussed that CTGF expression in hepatocytes (and other cell types) is efficiently stimulated not only by exogenous TGF-b but to a similar extent also by exogenous Activin A, with both cytokines sharing a common intracellular signaling pathway, involving phosphorylation of Smad2/3 (Gressner et al., 2002; Miyazawa et al., 2002; Pangas and Woodruff, 2000). Since Activin A occurs both intra- and extracellularly (Roth-Eichhorn et al., 1998), and even in the blood (Re and Cook, 2007, 2008), auto- and paracrine pathways might be relevant. And just as for TGF-b itself, intracrine signaling loops leading to target gene (i.e., CTGF) induction have also been described for Activin A in liver parenchymal (and other) cells (Gressner et al., 2008a; Miyazawa et al., 2002). However, different from TGF-b, increasing intracellular availability of Activin A in times of hepatocellular stress in vitro is not the result of its cleavage from a latent complex, but of a stimulation of its de novo synthesis (Gressner et al., 2008a). Also, the positive immunostaining of Activin A in hepatocytes of livers injured for various times with intraperitoneal applications of CCl4 is due to an increase in newly synthesized Activin A under injurious conditions (Gressner et al., 2008b). The conclusion of a stimulatory intracrine signaling mechanism also of Activin A, which was elaborated by our group, is based on the failure of the known extracellularly acting Activin A inhibitors follistatin, the physiological antagonist of Activin A, and neutralizing a-Activin A antibody to downregulate basal CTGF expression in hepatocytes, which was in contrast to the intracellularly acting TGF-b type I (Alk4/5) receptor serine– threonine kinase inhibitor SB431542 (Gressner et al., 2008a). Also, gene silencing of inhibin bA, that is, the bA subunit of Activin A, had a strong inhibitory effect on CTGF expression (Gressner et al., 2008a). Thus, not
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Figure 4.8 Extracellular activation of latent TGF-b. Activin A is newly synthesized in hepatocytes within the first 24 h after acute cellular stress in culture or in vivo toxicity, and leads to an activation of the kinase domain of the Alk4-receptor. By doing so, it triggers Smad2- and Smad3-phosphorylation leading to Activin A target gene (e.g., CTGF) expression. SARA, Smad anchor for receptor activation.
only the induction of CTGF synthesis by Activin A per se was shown but also that the cytokine does not necessarily require contact with the extracellular space in order to initiate target gene activation. However, in contrast to the stimulatory intracrine signaling pathway triggered by TGF-b, intracellular signaling of Activin A seems to involve both Smad2 and Smad3 phosphorylation and not only Smad2 phosphorylation (Gressner et al., 2008a) (Fig. 4.8).
VIII. Conclusion Under consideration of the current literature, it may be summarized that in conditions of in situ and cultural cellular stress, the cell seems to be exposed to a balance between autocrine BMP signaling and intracrine
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Activin A or TGF-b signaling following the release of the latter cytokine from intracellular latency. Activin A shares common Smad signaling pathways with the master cytokine TGF-b, leading to specific target gene (e.g., CTGF) expression. Our data raise the question on the physiologic role and pathophysiologic meaning of early intracrine Activin A and TGF-b signaling in parenchymal cells such as hepatocytes (and other cells). The pleiotropic functions, which are ascribed to both cytokines in the liver (ten Dijke et al., 2000) such as in other organs, might be dependent, at least partially, on the functions of its transcriptional targets such as CTGF. Up to now, the action of Activin A on the regulation of parenchymal cell proliferation, induction of apoptosis, development of fibrosis, tubulogenesis of endothelial cells, restoration of tissue architecture during regeneration, and, finally, even on carcinogenesis in several organs are discussed independently or in context with an induction of CTGF expression (Kreidl et al., 2009). Up to now, most of the proposed activities of CTGF are related to the function of TGF-b leading to its designation as “matricellular and oncogenic signaling modulator” (RothEichhorn et al., 1998). Interestingly, CTGF is suggested to modulate the balance of cell signaling by TGF-b and BMPs, respectively (Roth et al., 1998). An activation of TGF-b and inhibition of BMP signaling by affecting their binding to the respective receptors was clearly shown (Roth et al., 1998). Furthermore, it should be noted that CTGF is suggested as a mitogenic signal factor required for G0/G1 transition (Tan et al., 2004; Wiley et al., 2003). Such a role would convincingly explain the prometastatic/oncogenic and profibrogenic effect of CTGF, and also of TGF-b, which is supported by recent studies, in which knockdown of CTGF by siRNA leads to substantial attenuation of experimental liver fibrosis (Gao et al., 1996; Gressner and Wulbrand, 1997). One might speculate on a similar action of CTGF on Activin A signaling, but its causal and timely relation to Activin A under special consideration of intracrine signaling as initial event following cellular injury still remains to be analyzed.
REFERENCES Aaronson, S. A., Igarashi, H., Rao, C. D., Finzi, E., Fleming, T. P., Segatto, O., and Robbins, K. C. (1986). Role of genes for normal growth factors in human malignancy. Princess Takamatsu Symp. 17, 95–108. Arora, A. S., de Groen, P., Emori, Y., and Gores, G. J. (1996). A cascade of degradative hydrolase activity contributes to hepatocyte necrosis during anoxia. Am. J. Physiol. 270, G238–G245. Bejcek, B. E., Li, D. Y., and Deuel, T. F. (1989). Transformation by v-sis occurs by an internal autoactivation mechanism. Science 245, 1496–1499. Browder, T. M., Dunbar, C. E., and Nienhuis, A. W. (1989). Private and public autocrine loops in neoplastic cells. Cancer Cells 1, 9–17.
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Chen, D., Zhao, M., and Mundy, G. R. (2004). Bone morphogenetic proteins. Growth Factors 22, 233–241. Dallas, S. L., Park-Snyder, S., Miyazono, K., Twardzik, D., Mundy, G. R., and Bonewald, L. F. (1994). Characterization and autoregulation of latent transforming growth factor beta (TGF beta) complexes in osteoblast-like cell lines. Production of a latent complex lacking the latent TGF beta-binding protein. J. Biol. Chem. 269, 6815–6821. De Bleser, P. J., Niki, T., Rogiers, V., and Geerts, A. (1997). Transforming growth factorbeta gene expression in normal and fibrotic rat liver. J. Hepatol. 26, 886–893. Dowdy, S. C., Mariani, A., and Janknecht, R. (2003). HER2/Neu- and TAK1-mediated up-regulation of the transforming growth factor beta inhibitor Smad7 via the ETS protein ER81. J. Biol. Chem. 278, 44377–44384. Eijken, M., Swagemakers, S., Koedam, M., Steenbergen, C., Derkx, P., Uitterlinden, A. G., van der Spek, P. J., et al. (2007). The activin A-follistatin system: Potent regulator of human extracellular matrix mineralization. FASEB J. 21, 2949–2960. Fan, J., Shen, H., Sun, Y., Li, P., Burczynski, F., Namaka, M., and Gong, Y. (2006). Bone morphogenetic protein 4 mediates bile duct ligation induced liver fibrosis through activation of Smad1 and ERK1/2 in rat hepatic stellate cells. J. Cell. Physiol. 207, 499–505. Fernandez, T., Amoroso, S., Sharpe, S., Jones, G. M., Bliskovski, V., Kovalchuk, A., Wakefield, L. M., et al. (2002). Disruption of transforming growth factor beta signaling by a novel ligand-dependent mechanism. J. Exp. Med. 195, 1247–1255. Fleming, T. P., Matsui, T., Molloy, C. J., Robbins, K. C., and Aaronson, S. A. (1989). Autocrine mechanism for v-sis transformation requires cell surface localization of internally activated growth factor receptors. Proc. Natl. Acad. Sci. USA 86, 8063–8067. Gao, C., Gressner, G., Zoremba, M., and Gressner, A. M. (1996). Transforming growth factor beta (TGF-beta) expression in isolated and cultured rat hepatocytes. J. Cell. Physiol. 167, 394–405. Gkretsi, V., Mars, W. M., Bowen, W. C., Barua, L., Yang, Y., Guo, L., St-Arnaud, R., et al. (2007). Loss of integrin linked kinase from mouse hepatocytes in vitro and in vivo results in apoptosis and hepatitis. Hepatology 45, 1025–1034. Gleizes, P. E., Beavis, R. C., Mazzieri, R., Shen, B., and Rifkin, D. B. (1996). Identification and characterization of an eight-cysteine repeat of the latent transforming growth factorbeta binding protein-1 that mediates bonding to the latent transforming growth factorbeta1. J. Biol. Chem. 271, 29891–29896. Gonzalez-Gaitan, M. (2008). The garden of forking paths: Recycling, signaling, and degradation. Dev. Cell 15, 172–174. Gressner, O. A., and Gressner, A. M. (2008). Connective tissue growth factor: A fibrogenic master switch in fibrotic liver diseases. Liver Int. 28, 1065–1079. Gressner, A. M., and Wulbrand, U. (1997). Variation of immunocytochemical expression of transforming growth factor (TGF)-beta in hepatocytes in culture and liver slices. Cell Tissue Res. 287, 143–152. Gressner, A. M., Lahme, B., and Roth, S. (1997). Attenuation of TGF-beta-induced apoptosis in primary cultures of hepatocytes by calpain inhibitors. Biochem. Biophys. Res. Commun. 231, 457–462. Gressner, A. M., Weiskirchen, R., Breitkopf, K., and Dooley, S. (2002). Roles of TGF-beta in hepatic fibrosis. Front. Biosci. 7, d793–d807. Gressner, O. A., Lahme, B., Demirci, I., Gressner, A. M., and Weiskirchen, R. (2007). Differential effects of TGF-beta on connective tissue growth factor (CTGF/CCN2) expression in hepatic stellate cells and hepatocytes. J. Hepatol. 47, 699–710.
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Gressner, O. A., Lahme, B., Siluschek, M., Rehbein, K., Weiskirchen, R., and Gressner, A. M. (2008a). Intracrine signalling of activin A in hepatocytes upregulates connective tissue growth factor (CTGF/CCN2) expression. Liver Int. 28, 1207–1216. Gressner, O. A., Lahme, B., Siluschek, M., Rehbein, K., Herrmann, J., Weiskirchen, R., and Gressner, A. M. (2008b). Activation of TGF-beta within cultured hepatocytes and in liver injury leads to intracrine signaling with expression of connective tissue growth factor. J. Cell. Mol. Med. 12(6B), 2717–2730. Gressner, O. A., Lahme, B., Siluschek, M., Rehbein, K., Weiskirchen, R., and Gressner, A. M. (2009). Connective tissue growth factor is a Smad2 regulated amplifier of transforming growth factor-beta actions in hepatocytes—But without modulating bone morphogenetic protein 7 signalling. Hepatology 10.1002/hep. 22850. Grotendorst, G. R. (1997). Connective tissue growth factor: A mediator of TGF-beta action on fibroblasts. Cytokine Growth Factor Rev. 8, 171–179. Inagaki, Y., and Okazaki, I. (2007). Emerging insights into Transforming growth factor beta Smad signal in hepatic fibrogenesis. Gut 56, 284–292. Inman, G. J., Nicolas, F. J., Callahan, J. F., Harling, J. D., Gaster, L. M., Reith, A. D., Laping, N. J., et al. (2002). SB-431542 is a potent and specific inhibitor of transforming growth factor-beta superfamily type I activin receptor-like kinase (ALK) receptors ALK4, ALK5, and ALK7. Mol. Pharmacol. 62, 65–74. Inomata, M., Hayashi, M., Ohno-Iwashita, Y., Tsubuki, S., Saido, T. C., and Kawashima, S. (1996). Involvement of calpain in integrin-mediated signal transduction. Arch. Biochem. Biophys. 328, 129–134. Jachimczak, P., Hessdorfer, B., Fabel-Schulte, K., Wismeth, C., Brysch, W., Schlingensiepen, K. H., Bauer, A., et al. (1996). Transforming growth factor-betamediated autocrine growth regulation of gliomas as detected with phosphorothioate antisense oligonucleotides. Int. J. Cancer 65, 332–337. Koli, K., Saharinen, J., Hyytiainen, M., Penttinen, C., and Keski-Oja, J. (2001). Latency, activation, and binding proteins of TGF-beta. Microsc. Res. Tech. 52, 354–362. ¨ ztu¨rk, D., Metzner, T., Berger, W., and Grusch, M. (2009). Activins and Kreidl, E., O follistatins: Emerging roles in liver physiologyand cancer. World J. Gastroenterol. 1, 17–27. Leask, A., and Abraham, D. J. (2006). All in the CCN family: Essential matricellular signaling modulators emerge from the bunker. J. Cell Sci. 119, 4803–4810. Leask, A., Sa, S., Holmes, A., Shiwen, X., Black, C. M., and Abraham, D. J. (2001). The control of ccn2 (ctgf) gene expression in normal and scleroderma fibroblasts. Mol. Pathol. 54, 180–183. Leask, A., Holmes, A., Black, C. M., and Abraham, D. J. (2003). Connective tissue growth factor gene regulation. Requirements for its induction by transforming growth factorbeta 2 in fibroblasts. J. Biol. Chem. 278, 13008–13015. Limaye, P. B., Apte, U. M., Shankar, K., Bucci, T. J., Warbritton, A., and Mehendale, H. M. (2003). Calpain released from dying hepatocytes mediates progression of acute liver injury induced by model hepatotoxicants. Toxicol. Appl. Pharmacol. 191, 211–226. Massague, J. (1998). TGF-beta signal transduction. Annu. Rev. Biochem. 67, 753–791. Mehendale, H. M., and Limaye, P. B. (2005). Calpain: A death protein that mediates progression of liver injury. Trends Pharmacol. Sci. 26, 232–236. Miyazawa, K., Shinozaki, M., Hara, T., Furuya, T., and Miyazono, K. (2002). Two major Smad pathways in TGF-beta superfamily signalling. Genes Cells 7, 1191–1204. Moussad, E. E., and Brigstock, D. R. (2000). Connective tissue growth factor: What’s in a name? Mol. Genet. Metab. 71, 276–292. Nohe, A., Keating, E., Knaus, P., and Petersen, N. O. (2004). Signal transduction of bone morphogenetic protein receptors. Cell. Signal. 16, 291–299.
Intracrine Signaling of Activin A and TGF-b
77
Pangas, S. A., and Woodruff, T. K. (2000). Activin signal transduction pathways. Trends Endocrinol. Metab. 11, 309–314. Re, R. N., and Cook, J. L. (2006). The intracrine hypothesis: An update. Regul. Pept. 133, 1–9. Re, R. N., and Cook, J. L. (2007). Potential therapeutic implications of intracrine angiogenesis. Med. Hypotheses 69, 414–421. Re, R. N., and Cook, J. L. (2008). The basis of an intracrine pharmacology. J. Clin. Pharmacol. 48, 344–350. Roberts, A. B., Anzano, M. A., Lamb, L. C., Smith, J. M., and Sporn, M. B. (1981). New class of transforming growth factors potentiated by epidermal growth factor: Isolation from non-neoplastic tissues. Proc. Natl. Acad. Sci. USA 78, 5339–5343. Roberts, A. B., Kim, S. J., and Sporn, M. B. (1991). Is there a common pathway mediating growth inhibition by TGF-beta and the retinoblastoma gene product? Cancer Cells 3, 19–21. Roth, S., Schurek, J., and Gressner, A. M. (1997). Expression and release of the latent transforming growth factor beta binding protein by hepatocytes from rat liver. Hepatology 25, 1398–1405. Roth, S., Michel, K., and Gressner, A. M. (1998). (Latent) transforming growth factor beta in liver parenchymal cells, its injury-dependent release, and paracrine effects on rat hepatic stellate cells. Hepatology 27, 1003–1012. Roth-Eichhorn, S., Kuhl, K., and Gressner, A. M. (1998). Subcellular localization of (latent) transforming growth factor beta and the latent TGF-beta binding protein in rat hepatocytes and hepatic stellate cells. Hepatology 28, 1588–1596. Saharinen, J., Taipale, J., and Keski-Oja, J. (1996). Association of the small latent transforming growth factor-beta with an eight cysteine repeat of its binding protein LTBP-1. EMBO J. 15, 245–253. Seyhan, H., Hamzavi, J., Wiercinska, E., Gressner, A. M., Mertens, P. R., Kopp, J., Horch, R. E., et al. (2006). Liver fibrogenesis due to cholestasis is associated with increased Smad7 expression and Smad3 signaling. J. Cell. Mol. Med. 10, 922–932. Tan, C. M., Brady, A. E., Nickols, H. H., Wang, Q., and Limbird, L. E. (2004). Membrane trafficking of G protein-coupled receptors. Annu. Rev. Pharmacol. Toxicol. 44, 559–609. ten Dijke, P., Miyazono, K., and Heldin, C. H. (2000). Signaling inputs converge on nuclear effectors in TGF-beta signaling. Trends Biochem. Sci. 25, 64–70. Tsukazaki, T., Chiang, T. A., Davison, A. F., Attisano, L., and Wrana, J. L. (1998). SARA, a FYVE domain protein that recruits Smad2 to the TGFbeta receptor. Cell 95, 779–791. Wells, R. G., Yankelev, H., Lin, H. Y., and Lodish, H. F. (1997). Biosynthesis of the type I and type II TGF-beta receptors. Implications for complex formation. J. Biol. Chem. 272, 11444–11451. Whitman, M. (1997). Signal transduction. Feedback from inhibitory SMADs. Nature 389, 549–551. Wiley, H. S., Shvartsman, S. Y., and Lauffenburger, D. A. (2003). Computational modeling of the EGF-receptor system: A paradigm for systems biology. Trends Cell Biol. 13, 43–50. Wu, L., and Derynck, R. (2009). Essential role of TGF-beta signaling in glucose-induced cell hypertrophy. Dev. Cell 17, 35–48.
C H A P T E R
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Negative Regulation of Activin Signal Transduction Sun-Cheol Choi1 and Jin-Kwan Han Contents 80 82 82 83 85 85 85 87 87 88 89 89 90 91 91 92 93 94 94 97 98 98
I. Introduction II. Extracellular Antagonists A. Inhibin B. Follistatin and follistatin-related gene III. Negative Regulation of Activin Receptor Function A. BMP and activin membrane-bound inhibitor B. Cripto C. Regulatory subunits of PP2A (Ba and Bd) D. Activin receptor-interacting protein 1 and 2 E. Dok-PTB containing protein F. Dapper2 IV. Inhibitory Smads V. Interfering with Smad Function A. PPM1A B. Ectodermin/TIF1g C. Linker phosphorylation of Smads D. Erbin E. Transmembrane prostate androgen-induced RNA VI. Negative Regulation of Gene Transcription VII. Conclusions Acknowledgments References
Abstract Activin is a member of the transforming growth factor b (TGFb) superfamily. While it was originally isolated as a gonadal factor to regulate secretion of follicle-stimulating hormone (FSH) from the pituitary, it also has nonreproductive roles in immune responses, metabolism, tumorigenesis, and stem cell Division of Molecular and Life Sciences, Pohang University of Science and Technology, Hyoja-dong, Nam-gu, Pohang, Kyungbuk, Republic of Korea Current address: Department of Medicine, Graduate School, University of Ulsan, Pungnap-Dong, Songpa-Gu, Seoul, Republic of Korea
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Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00005-6
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2011 Elsevier Inc. All rights reserved.
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differentiation. Activin signaling is initiated by ligand-induced formation of a heteromeric complex of type I and type II transmembrane serine/threonine kinase receptors. The activated activin receptors phosphorylate the receptorregulated Smads, Smad2 and Smad3, which subsequently form a complex with the common mediator, Smad4, and translocate into the nucleus for the transcriptional regulation of specific target genes in cooperation with DNA-binding cofactors and transcriptional coactivators. Activin signaling is controlled both extracellularly and intracellularly by diverse mechanisms to fine tune its duration and strength. This chapter summarizes current understanding of how activin signaling pathway is negatively regulated inside and outside the cells. ß 2011 Elsevier Inc.
I. Introduction Activin belongs to the transforming growth factor b (TGFb) superfamily, which includes TGFb, activin, nodal, bone morphogenetic proteins (BMPs), myostatin, and others. These secreted cytokines regulate a variety of cellular responses such as differentiation, proliferation, migration, adhesion, and apoptosis (ten Dijke and Hill, 2004). Activin was first discovered as a gonadal regulator that activates release of follicle-stimulating hormone (FSH) from the anterior pituitary. Besides its reproductive roles, activin has been shown to be implicated in metabolism, immune response, differentiation of embryonic stem cell, tumorigenesis, and brain function (Welt et al., 2002; Xia and Schneyer, 2009). Activin is a dimer composed of two b subunits. bA and bB subunits form homodimeric activin A (bA–bA) or activin B (bB–bB) and heterodimeric activin AB (bA–bB). Activin signals through a heterotetrameric complex of specific type I and type II transmembrane serine/threonine kinase receptors (Fig. 5.1). Two subtypes of the type II activin receptor, ActRIIA and ActRIIB, have been identified. In addition, ALK4 (also known as ActRIB) and ALK7 (ACVR1C) work as the type I receptor for activin. These activin receptors are shared by other members of TGFb family such as nodal and growth and differentiation factor 11 (GDF11). Activin binds directly to the constitutively active type II receptor, which then recruits and phosphorylates the type I receptor in the unique Gly-Ser (GS) domain near the juxtamembrane region. The activated type I receptor undergoes a conformational change, thereby facilitating the docking of specific receptor-regulated Smad (R-Smad) proteins and activating them through phosphorylation at the C-terminal serine residues. While Smad2 and Smad3 are phosphorylated by TGFb, activin and nodal type I receptors, Smad1, Smad5, and Smad8, are activated by BMP type I receptor. The R-Smads contain two conserved domains, MH1 and MH2, which are separated by a linker region. The N-terminal MH1 domain has DNA-binding activity, whereas the
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Figure 5.1 Negative regulators of activin signaling. Activin binds to a heteromeric receptor complex and induces type II receptor phosphorylation of type I receptor. Extracellular inhibitors, including inhibin, follistatin, and follistatin-related genes (FLRGs), and membrane proteins such as Cripto and BAMBI prevent the access of activin ligand to its receptors and/or the formation of the functional activin receptor complex. The activated type I receptor recruits and phosphorylates Smad2/3, which subsequently form a complex with Smad4 and go to the nucleus to regulate gene transcription. It has been shown that clathrin-dependent endocytosis promotes receptor activation of Smad2/3. SARA facilitates receptor recruitment and activation of Smad2/3 on early endosome, which is antagonized by a transmembrane protein, TMEPAI. However, the receptors internalized through a caveolin-dependent route undergo Smad7/Smurf2 complex-mediated degradation. Smad7-interacting proteins such as FKBP12 and salt-inducible kinase (SIK) promote receptor degradation, whereas Rap2 GTPase and Hsp90 contribute to receptor stabilization. In the absence of ligand, the homodimeric receptor complexes are internalized and then recycle to the cell surface through a Rab11-mediated pathway. Rap2 promotes receptor recycling, which is counteracted by Smad7. Receptor-associating proteins including ARIPs, Dpcp, Dapper2 and regulatory subunits of phosphatase PP2A, PPP2R2A/2D affect the stability, localization, and activity of receptors and/or receptor recruitment of Smad2/3. A cytoplasmic protein, Erbin interferes with the association of the activated Smad2/3 and Smad4.
C-terminal MH2 domain mediates protein–protein interaction for nuclear translocation and transcriptional regulation (Schmierer and Hill, 2007; Welt et al., 2002). The recruitment of R-Smads to the receptor complex is facilitated by scaffolding proteins such as Smad anchor for receptor activation (SARA) and Hgs (Hrs), which are FYVE domain proteins and cooperate
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synergistically in activin receptor-mediated signaling (Miura et al., 2000; Tsukazaki et al., 1998). Activated R-Smads form a complex with common Smad4 (Co-Smad), and this complex translocates to the nucleus to regulate the transcription of specific genes. Once there, Smad complexes bind to the promoter region of target genes with transcriptional factors such as FoxH1 and Mixer as well as transcriptional coactivator including p300 and CBP (Schmierer and Hill, 2007; ten Dijke and Hill, 2004). While activin signaling appears simply linear as described above, the complicated regulatory mechanisms, positive and negative, are actually involved at various steps of the signaling cascade in fine-tuning its duration and intensity. Here, we review these control mechanisms, mainly focusing on how activin signaling is negatively regulated both outside and inside the cells.
II. Extracellular Antagonists As a secreted protein, activin functions in an endocrine, autocrine, or paracrine manner (Welt et al., 2002). Thus, its availability for binding to the receptors is limited by extracellular activin-binding proteins such as inhibin and follistatin (Fig. 5.1).
A. Inhibin Inhibin was initially identified as a gonadal factor which plays negative roles in the secretion and synthesis of FSH by the pituitary gland and was later shown to be a strong antagonist of activin responses in many tissues (Bilezikjian et al., 2006; Gray et al., 2002). A common a-subunit is linked to one of the two activin b-subunits (bA and bB) by disulfide bonds to form the heterodimers, inhibin A (abA) and inhibin B (abB). Inhibin interacts preferentially with the type II activin receptor, ActRII, and is unable to block the activation of a reporter gene in response to a constitutively active type I activin receptor (Martens et al., 1997), suggesting its function at the level of the activin–receptor complex. It has been shown that inhibin antagonism of activin signaling is achieved by its displacement of activin from ActRIIs. While inhibin and activin share the same binding site on ActRII (Gray et al., 2000), the binding affinity of inhibin for the receptor is about 10-fold lower than that of activin (Mathews and Vale, 1991), which is inconsistent with a model of simple competition between activin and inhibin for binding to ActRII. Importantly, even molar excesses of inhibin fail to counteract activin signaling in some cell types such as corticotropes (Lebrun and Vale, 1997; Lewis et al., 2000), indicative of the requirement of additional components for the inhibitory effects of inhibin. It has been demonstrated that betaglycan, a membrane-anchored proteoglycan and
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originally identified as the type III TGFb receptor, can act as an inhibin coreceptor with ActRII (Lewis et al., 2000). Betaglycan binds inhibin with high affinity and increases the binding of inhibin to cell membranes when coexpressed with ActRII. It confers inhibin responsiveness to cells such as corticotrope AtT20 cells that otherwise are insensitive to inhibin action. Furthermore, the functional disruption of betaglycan through RNA interference-mediated knockdown and immunoneutralization impairs the ability of inhibin to antagonize activin-stimulated FSH secretion and reporter gene activation in gonadotropes (Escalona et al., 2009; Wiater et al., 2009), suggesting its essential role for the high potency of inhibin antagonism of activin responses in these cells. Cross-linking experiments have revealed that inhibin forms a complex with betaglycan and ActRII but not with the type I activin receptor, ALK4 (Lewis et al., 2000). Thus, these results suggest a model in which betaglycan facilitates the binding of inhibin to ActRII to form a ternary complex without recruiting ALK4 and prevents access of activin to the receptor, thereby leading to a blockade of activin signaling.
B. Follistatin and follistatin-related gene Follistatin was originally isolated as a component of follicular fluids with the ability to suppress FSH secretion from pituitary cells (Welt et al., 2002). It is a monomeric glycosylated polypeptide that binds and neutralizes activin with high affinity, and the binding is nearly irreversible due to its slow dissociation rate (Schneyer et al., 1994). Follistatin is composed of a 63-residue N-terminal segment, followed by three successive 73–75-residue follistatin (FS) domains, with each containing ten cysteine amino acids (Xia and Schneyer, 2009). The N-terminal domain and FS domains 1 and 2, but not C-terminal FS domain 3, were found to be critical for activin binding and inhibiting activin responses (Keutmann et al., 2004; Sidis et al., 2001). Moreover, rearrangement or duplication of the FS domains diminished activin binding and neutralization (Keutmann et al., 2004), suggesting that the number and sequential order of the FS domains are essential for the full activity of follistatin. Structural analysis of the follistatin–activin A complex revealed that two follistatin molecules encircle the activin dimer, covering a large proportion of its residues and blocking both type I and type II receptor binding sites (Thompson et al., 2005), which leads to prevention of activin from interacting with the receptor complex and abrogation of activin signaling. Two variants of follistatin are generated through alternative splicing at the C-terminus. The shorter FS288 isoform terminates after the third FS domain, and the heparin-binding sites in its FS domain 1 render this isoform capable of binding heparin-sulfated proteoglycans on the cell surface with high affinity and potentially impeding action of autocrine-acting activin (Welt et al., 2002). This FS isoform-sequestered activin at the cell
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surface might undergo accelerated endocytic internalization and subsequent degradation (Hashimoto et al., 1997). The longer isoform, FS315, is extended to 315 residues by an acidic C-terminal extension encoded by an extra exon and is localized predominantly in the circulation (Harrison et al., 2005; Welt et al., 2002). Consistently, it has a reduced affinity for heparin possibly due to the masking of the heparin-binding site by the acidic C-terminal extension. Proteolytic processing of the C-terminal acidic tail of FS315 produces the intermediate isoform, FS303, which is found primarily in gonadal fluids and binds partially cell surface heparin-sulfated proteoglycans (Welt et al., 2002). Follistatin-related gene (FLRG), which is also known as follistatinrelated protein (FLRP) or follistatin-like 3 (FSTL3), is a follistatin-domain containing protein that binds activin with high affinity and low reversibility (Tsuchida et al., 2000). While both FS288 and FS315 have three FS domains, FLRG lacks the third FS domain. The N-terminal domains are significantly divergent between FS and FLRG (24% sequence identity), whereas the sequences of FS domains 1 and 2 are relatively well conserved. FLRG also lacks a consensus heparin-binding motif and does not bind cell surface heparin-sulfated proteoglycans, which makes it act as a circulating activin-binding protein (Schneyer et al., 2001). Thus, it efficiently inhibits the transcriptional responses stimulated by activin in a paracrine/endocrine fashion but not FSH secretion in response to endogenously produced activin (autocrine-acting activin) in cultured pituitary cells (Schneyer et al., 2001), suggesting that the structural differences between FS and FLRG might confer distinct bioactivities. A recently determined structure of FLRG in complex with activin A has shown that two FLRG molecules surround the ligand, blocking all receptor binding sites as observed in the FS–activin A complex (Stamler et al., 2008). However, the N-terminal domain of FLRG adopts a more compact conformation in contacting activin than that of FS, possibly compensating for the lack of the third FS domain in FLRG. In addition, exchange of the N-terminal domain in FLRG with that of FS reduces considerably its affinity for activin (Stamler et al., 2008), implying that the domains in FS and FLRG form a unique but significant contact at the ligand interface. Tissue distribution, subcellular localization, and intracellular transport pattern of FS and FLRG also suggest that they may not be complete functional homologs. Follistatin is highly expressed in ovary and pituitary, whereas FLRG is distributed predominantly in testis, placenta, heart, and pancreas (Schneyer et al., 2001). Unlike follistatin, FLRG is localized in the nucleus though it is also secreted (Tortoriello et al., 2001). However, FLRG is the slowest to be secreted compared to FS288 and FS315 (Saito et al., 2005). The nuclear FLRG is also N-glycosylated but not to the same degree as secreted FLRG. These results indicate that FS and FLRG may have distinct intracellular and/or extracellular functions.
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III. Negative Regulation of Activin Receptor Function Membrane or cytoplasmic proteins interfere with activin receptormediated signaling by negatively affecting the assembly of functional receptors, the stability, activity, and localization of receptors or receptor recruitment of R-Smads (Fig. 5.1).
A. BMP and activin membrane-bound inhibitor The transmembrane glycoprotein BAMBI (BMP and activin membranebound inhibitor) is closely related to the type I receptors of the TGFb superfamily in the extracellular domain but lacks an intracellular kinase domain (Grotewold et al., 2001; Onichtchouk et al., 1999). It is tightly coexpressed with BMP4 during early embryo development of Xenopus, mouse, and zebrafish (Grotewold et al., 2001; Onichtchouk et al., 1999; Tsang et al., 2000), and its expression is induced by BMP4, TGFb, and Wnt signaling (Sekiya et al., 2004a,b). Its human homolog, nma, is downregulated in metastatic melanoma cell lines, and its elevated expression suppresses TGFb-mediated growth inhibition in colorectal and hepatocellular carcinomas (Degen et al., 1996; Sekiya et al., 2004a). BAMBI acts as a pseudoreceptor to stably interact with both type I and type II receptors in a ligand-independent manner, inhibiting the signaling by BMP, activin, and TGFb ligands (Onichtchouk et al., 1999). Recently, it has been shown that BAMBI exploits an alternative mechanism to block TGFb signaling in which it forms a ternary complex with the TGFb type I receptor, ALK5, and an inhibitory Smad7, and impedes the association of ALK5 and Smad3, thereby blocking Smad3 activation (Yan et al., 2009). Thus, it has been suggested that BAMBI could utilize dual mechanisms to attenuate TGFb signaling, and these two mechanisms might work cooperatively. Further experiments are required to confirm that BAMBI could also inhibit activin signaling through the second mechanism involving Smad7 function.
B. Cripto Cripto is a member of the epidermal growth factor (EGF)-Cripto-FRL1Cryptic (CFC) protein family and has activity both as a glycosylphosphatidylinositol (GPI)-anchored membrane protein and a soluble factor (Strizzi et al., 2005). It is expressed at high levels in the human breast, lung, colon, and the ovarian carcinomas, and its overexpression promotes a variety of tumorigenic properties such as cell proliferation, migration, and epithelial-to-mesenchymal transition (EMT) (Bianco et al., 2004).
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In the developing embryo, Cripto also functions as a coreceptor for TGFb ligands, including Nodal, Vg1, and GDF-1 (Cheng et al., 2003; Schier, 2003). Nodal ligand signals through an ActRII/IIB–ALK4–Cripto receptor complex to activate the cytoplasmic Smad2/3 pathway and nuclear gene expression. Cripto directly interacts with ALK4, thereby enabling the nodal ligand to bind the ActRII/IIB–ALK4 complex (Yeo and Whitman, 2001). Mutagenesis experiments have revealed that Cripto binds to the nodal ligand via its EGF-like domain and ALK4 via its CFC domain (Yeo and Whitman, 2001). Genetic and embryological studies in zebrafish, mice, and frog have shown that Cripto-dependent nodal signaling is essential for the patterning of the anterior–posterior axis, formation of mesoderm and endoderm, and establishment of the left–right asymmetric axis (Hamada et al., 2002; Schier, 2003). In contrast to its requirement for nodal signaling, Cripto inhibits activin signaling. In one study, Cripto has been shown to interact directly with activin B but not with activin A ligand (Adkins et al., 2003). It has been proposed that Cripto abrogates activin B signaling by binding and sequestering the ligand from the type II activin receptor and/or forming nonfunctional complexes with the type I receptor, ALK4. Gray et al. (2003) have also observed the negative effect of Cripto on activin signaling, but they did not detect activin binding to Cripto in the absence of ActRII/IIB, suggesting the requirement of the type II receptors for activin to bind Cripto. They presented additional evidence that Cripto in complex with the activin–ActRII/IIB could impede the subsequent recruitment of ALK4, blocking downstream signaling events. Although these two groups reported distinct inhibitory complexes, a common mechanism for Cripto to block activin signaling seems to involve its competitive inhibition of receptor assembly. Interestingly, another novel mechanism was recently proposed that Cripto acts as a noncompetitive activin antagonist, forming functional analogous receptor complex with activin and nodal (Kelber et al., 2008). In this model, Cripto just reduces activin signaling capacity without affecting its affinity for ActRII/IIB. Thus, activin and nodal induce similar maximal signaling responses in the presence of Cripto, which are considerably lower than that elicited by activin in the absence of Cripto. This mechanism seems to address the apparently contradictory effects of Cripto on activin and nodal signaling which employ the same type I and type II receptors. The inhibitory effects of Cripto on activin signaling appear to involve a cell surface glucose-regulated protein 78 (GRP78), which is an endoplasmic reticulum (ER) chaperone in the heat-shock protein 70 family and binds to Cripto (Kelber et al., 2009). shRNA-mediated knockdown of Cripto and GRP78 enhances activin signaling responses such as Smad2 phosphorylation and activation of a reporter gene to a greater extent than knockdown of either protein alone, indicating their functional cooperation. Furthermore,
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a GRP78 blocking antibody that disrupts the interaction of Cripto and GRP78 inhibited Cripto-dependent effects on activin signaling. Thus, GRP78 functions as a mediator of Cripto signaling to modulate activin responses.
C. Regulatory subunits of PP2A (Ba and Bd) Ba (PPP2R2A) and Bd (PPP2R2D) are highly related members of the B family of regulatory subunits of the multimeric serine/threonine protein phosphatase PP2A ( Janssens and Goris, 2001). Knockdown of Ba or overexpression of Bd inhibits target gene expression and Smad2 phosphorylation induced by activin and nodal signals in Xenopus embryos and cultured cells, whereas knockdown of Bd exhibits opposing effects on these responses (Batut et al., 2008), suggesting that these two isoforms regulate activin/ nodal signaling in different ways, with Ba acting positively and Bd acting negatively. It has been documented that Ba functions to stabilize the basal levels of type I receptor with Bd restricting receptor activity (Batut et al., 2008). The exact mechanisms of their regulation of activin signaling await further analysis.
D. Activin receptor-interacting protein 1 and 2 Activin receptor-interacting proteins (ARIPs) were identified in a yeast two-hybrid screening using the cytoplasmic region of the ActRII, which harbors a consensus PDZ-binding motif, as a bait to search for the candidate molecules which would mediate the specific functions of the receptor (Matsuzaki et al., 2002; Shoji et al., 2000; Tsuchida et al., 2004). ARIP1 is abundant in the brain tissue and has two isoforms, ARIP1-long and ARIP1short, which might be generated by alternative splicing (Shoji et al., 2000). ARIP1-long contains one guanylate kinase domain in the N-terminal region, followed by two WW domains and five PDZ domains, whereas ARIP1-short lacks the guanylate kinase domain. Both isoforms interact with ActRIIA via their PDZ domains but not with ActRIIB, TGFb type II receptor, and BMP type II receptor. They also associate with Smad3, a mediator of activin signaling through the WW domains. Overexpression of ARIP1 suppresses activin or Smad3-induced transcriptional activation. Thus, it has been suggested that ARIP1 functions as a scaffolding protein to assemble activin receptors with a downstream mediator, Smad, regulating activin signaling in neuronal cells (Shoji et al., 2000). However, it remains to be investigated whether ARIP1 could inhibit activin signaling under physiological conditions. ARIP2 is a small cytoplasmic protein that has one PDZ domain and interacts with ActRIIA and ActRIIB via its PDZ domain (Matsuzaki et al., 2002). It also has two isoforms, ARIP2b and ARIP2c, which have a single
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PDZ domain but differ from ARIP2 in the C-terminal sequences (Liu et al., 2006). These isoforms interact specifically with ActRIIA but not with ActRIIB. While ARIP2 is detected in multiple mouse tissues, ARIP2b and 2c are highly expressed in the heart, liver, kidney, and the ovary (Liu et al., 2006). Overexpression of ARIP2 inhibits the activin-induced transcriptional responses and FSH secretion from pituitary cells and enhances cell proliferation in human breast carcinomas (Li et al., 2009; Liu et al., 2009; Matsuzaki et al., 2002). Conversely, RNAi-mediated knockdown of ARIP2 exerts opposing effects on these responses (Li et al., 2009), suggesting that ARIP2 could be involved in the negative regulation of activin signaling. Interestingly, unlike ARIP2, ARIP2b and 2c have stimulatory effects on the activin-induced transcriptional activity and FSH secretion from gonadotroph cells (Liu et al., 2006). Furthermore, ARIP2 and ARIP2b/2c antagonize each other in control of activin responses. Thus, it has been suggested that ARIP2 and its isoforms might have a role in shaping morphogenetic gradients and in fine-tuning activin signaling during tissue formation (Liu et al., 2006). Mechanically, expression of ARIP2 increases endocytosis of ActRII and reduces ActRIIA expression on cell membrane through the Ral/RalBP1-dependent pathway (Matsuzaki et al., 2002). ARIP2 is also involved in the intracellular translocation of ActRII via PDZ domain-mediated interaction. Further studies are necessary to decipher the molecular mechanisms by which the effects of ARIP2 on the trafficking of ActRII lead to its negative control of activin signaling.
E. Dok-PTB containing protein Dpcp, named after Dok-PTB containing protein, was initially identified in Xenopus embryo and has a phosphotyrosine-binding (PTB) domain, which mediates a protein–protein interaction (Cheong et al., 2009). It is localized in the cytoplasm and/or plasma membrane in cultured cells and Xenopus embryonic cells. Overexpression of Dpcp reduces gene expression induced by activin and nodal signals in vitro and in vivo, whereas its knockdown mediated by an antisense morpholino oligo augments this response, suggesting the negative role of Dpcp in regulation of activin/nodal signaling (Cheong et al., 2009). It has been demonstrated that Dpcp binds to the type I activin receptor, ALK4 via its PTB domain, thereby competitively interfering with the interaction of ALK4 and Smad2 and blocking the subsequent phosphorylation of Smad2. Dpcp expression is restricted to the animal hemisphere of Xenopus embryo, which is devoid of mesoderminducing signals such as activin and nodal signaling. Thus, it has been suggested that Dpcp plays critical roles in the correct positioning of primitive germ layers during early embryogenesis (Cheong et al., 2009).
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F. Dapper2 Dapper was first identified as a Disheveled-associated antagonist of Wnt signaling in Xenopus embryo and has two isoforms, Dapper1 and 2 (Cheyette et al., 2002). Dapper2 is localized in late endosomes and binds to both ALK4 and ALK5, the type I receptors for activin and TGFb signaling (Zhang et al., 2004). Overexpression of Dapper2, but not Dapper1, inhibits activin- or TGFb-dependent activation of reporter genes in mammalian cells and mesoderm formation in zebrafish (Su et al., 2007; Zhang et al., 2004). In contrast, siRNA knockdown of Dapper2 increases this transcriptional response. It has been demonstrated that Dapper2 promotes the degradation of ALK4 and ALK5 through the lysosomal pathway (Zhang et al., 2004). The molecular mechanism by which this occurs remains unknown.
IV. Inhibitory Smads Inhibitory Smads (I-Smads), Smad6 and Smad7 are transcriptionally induced by TGFb family ligands and play negative roles in these signaling pathways, thus establishing a negative feedback loop (Moustakas and Heldin, 2009; ten Dijke and Hill, 2004). While Smad7 inhibits activin, TGFb, and BMP pathways, Smad6 is more specific for BMP signaling. Both Smad6 and Smad7 have the C-terminal MH2 domain but are more divergent from the R-Smads and Co-Smad in the N-terminal and central regions. I-Smads can inhibit TGFb signaling by diverse mechanisms. Smad7 binds to type I receptor via its MH2 domain, thereby competitively blocking the phosphorylation of R-Smads and the heterocomplex formation of R-Smads and Co-Smad (Itoh and ten Dijke, 2007). Smad7 interacts constitutively with the HECT type of E3 ubiquitin ligases, Smurf1 and Smurf2, and recruits them to the activated type I receptor, which leads to the degradation of active receptors through proteasomal and lysosomal pathways (Kavsak et al., 2000). Smad7 itself is also degraded in this process. Smad7 also recruits a phosphatase complex of GADD34–PP1c to the activated TGFb type I receptor, ALK5, thus dephosphorylating and inactivating the receptor (Shi et al., 2004). Similarly, protein phosphatase 1a was shown to be recruited by Smad7 to another TGFb type I receptor, ALK1, in endothelial cells and to dephosphorylate ALK1 (Valdimarsdottir et al., 2006). It has also been shown that Smad7 inhibits TGFb signaling in the nucleus independently of type I receptors. It specifically binds to the Smadresponsive element via its MH2 domain, disrupting the formation of the functional Smad–DNA complex in response to TGFb signal (Zhang et al., 2007).
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The TGFb and activin receptors are constitutively internalized and recycled to the cell surface through a Rab11-dependent pathway (Choi et al., 2008; Mitchell et al., 2004). In the absence of ligand stimulation, a small GTPase Rap2 facilitates receptor recycling to the plasma membrane, thereby preventing the receptors from entering into a degradation pathway (Choi et al., 2008). This contributes to maintaining the levels of receptors on the cell surface and cell responsiveness to extracellular ligands. Smad7 has been shown to antagonize Rap2 activity, interfering with the recycling of receptors and trapping them in the cytoplasm (Choi et al., 2008). Upon ligand binding, the internalized receptors do not go back to the cell surface and then undergo Smad7–Smurf2-mediated degradation after a while, leading to the termination of signaling. In this process, Rap2 counteracts Smad7 function, delaying the receptor turnover and prolonging the duration of signaling (Fig. 5.1). It remains to be investigated how Rap2 and Smad7 antagonize each other. Several proteins have been documented to interact with Smad7 and regulate TGFb/activin signaling. FKBP12 is a cytoplasmic protein that binds to the immunosuppressant drugs, rapamycin and Tacrolimus (FK506). In the absence of ligands, FKBP12 binds to the GS motif of type I receptors whose serine residues are phosphorylated by type II receptors, preventing the leaky signals. Upon ligand stimulation, FKBP12 is transiently dissociated from the type I receptor and thereafter forms a complex with Smad7, Smurf1, and type I receptor, promoting the ubiquitination and degradation of the receptor to terminate the signaling (Yamaguchi et al., 2006). Salt-inducible kinase (SIK), a target gene of TGFb/Smad signaling, interacts and cooperates with Smad7, facilitating receptor downregulation (Kowanetz et al., 2008). Its depletion enhances TGFb-dependent gene responses, suggesting its negative roles for this signaling. Conversely, the 90-kDa heat-shock protein (Hsp90) regulates positively TGFb signaling as a TGFb-receptor-interacting protein. Inhibition of Hsp90 function blocks TGFb-induced transcription and Smad2/3 phosphorylation and results in the Smurf2-dependent degradation of TGFb receptors (Wrighton et al., 2008), indicating its role at the level of receptor for TGFb signaling. Overall, these findings suggest that a variety of regulators work together to fine tune Smad7 activity for the duration and strength of TGFb/activin signaling. However, the correlation among these regulatory proteins and their relevance for developmental processes remain to be further analyzed.
V. Interfering with Smad Function Posttranslational modifications such as phosphorylation and ubiquitination can regulate the activity, subcellular localization, and stability of Smads. In addition, these kinds of modifications and regulatory proteins have been shown to affect the formation of R-Smad and Smad4 complex.
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A. PPM1A The levels of type I receptor-phosphorylated R-Smads determine the duration and strength of TGFb/activin signaling. Thus, dephosphorylation of these phospho-R-Smads is a key regulatory mechanism to terminate signaling. Recently, PPM1A/PP2Ca was identified as a nuclear phosphatase to dephosphorylate the R-Smads (Lin et al., 2006). PPM1A directly dephosphorylates the C-terminally phosphorylated Smad2 and Smad3, leading to disrupted interaction of Smad4 and Smad2 or Smad3 and nuclear export of Smad2/3. Loss of PPM1A function enhances TGFb-induced antiproliferative and transcriptional responses, whereas its gain of function antagonizes dorsalizing activity of nodal or nodal-like signals in zebrafish. However, PPM1A is not specific for R-Smads; it has other substrates, including axin, phosphatidylinositol-3 kinase, and CDK6 (Schmierer and Hill, 2007).
B. Ectodermin/TIF1g Ectodermin/TIF1g functions as a RING-type E3 ubiquitin ligase in the nucleus to antagonize TGFb/activin and BMP signaling. It specifies ectoderm in Xenopus embryos by restricting the mesoderm-inducing activity of activin/nodal signals to the mesodermal region (Dupont et al., 2005). It also attenuates cell growth inhibition induced by TGFb signals in various human cells and mediates TGFb-dependent erythroid differentiation in hematopoietic stem cells (He et al., 2006). Recently, it has been shown that Ectodermin/TIF1g acts as a monoubiquitin ligase for Smad4. Ectodermin/TIF1g-mediated ubiquitination of Smad4 interferes with the formation of the Smad2/Smad4 or Smad3/Smad4 complex in the nucleus and then causes nuclear export of Smad4 (Dupont et al., 2009). This might contribute to turning off signaling or raising the thresholds of cell responsiveness to ligands. Interestingly, FAM/USP9x acts as a deubiquitinating enzyme for the exported Smad4 in the cytoplasm as opposed to the action of Ectodermin/TIF1g, thereby recycling Smad4 and recovering its competence to mediate TGFb signaling (Dupont et al., 2009). Thus, it has been suggested that the monoubiquitination/deubiquitination cycle of Smad4 is a way for cells to modulate their responsiveness to TGFb ligands. However, He et al. (2006) have proposed that Ectodermin/TIF1g competes with Smad4 for binding to the receptor-activated Smad2/3 to stimulate TGFbinduced differentiation response, but not the antiproliferative response, in hematopoietic stem cells. However, they did not observe Smad4 ubiquitination by Ectodermin/TIF1g. Thus, it seems likely that depending on the cellular contexts, Ectodermin/TIF1g functions in different ways.
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C. Linker phosphorylation of Smads In addition to C-terminal tail phosphorylation by type I receptor, R-Smads undergo agonist- or antagonist-induced phosphorylation in their linker sequences between the MH1 and MH2 domains (Fig. 5.2; Feng and Derynck, 2005). Antagonists such as fibroblast growth factor (FGF) and EGF stimulate Erk MAP kinases, which phosphorylate the linker regions of Smad1 and Smad2/3, thereby attenuating agonist-induced nuclear accumulation of these Smads and impairing Smad-dependent transcriptional responses (Kretzschmar et al., 1999). Thus, oncogenic activation of Ras, which leads to MAPK stimulation, suppresses the antiproliferative effects of
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Figure 5.2 Linker phosphorylation of Smads. Antagonists, including FGF and EGF, induce ERK MAP kinase phosphorylation of Smad2/3 in their linker regions, which leads to cytoplasmic retention and degradation of the Smads. In agonists-induced signaling, the C-terminally phosphorylated Smad2/3 form a complex with Smad4 and translocate into the nucleus where the activated R-Smads undergo linker phosphorylation mediated by cyclin-dependent kinases, CDK8 and CDK9. This modification functions to promote Smad-dependent transcription and mark the R-Smads for later degradation. After being disengaged from Smad4, the linker phosphorylated Smads are C-terminally dephosphorylated by PPM1A and exit into the cytoplasm. The exported Smad2/3 are recognized by an E3 ubiquitin ligase, Nedd4L, and subsequently subjected to polyubiquitination and turnover. SGK1, belonging to the PKB/Akt kinase family, phosphorylates Nedd4L, suppressing its activity.
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TGFb signaling in some cells, though the effects of Ras/MAPK signaling on nuclear translocation of Smads are different depending on cell types and conditions (Funaba et al., 2002). In Xenopus embryos, ectodermal cells express mesodermal genes with Smd2 nuclear accumulation in response to activin signaling, and this mesodermal competence is abruptly lost with a nuclear exclusion of Smad2, which is dependent on the phosphorylation of Erk/MAPK sites in the linker region by some kinases (Grimm and Gurdon, 2002). The mechanism by which MAPK phosphorylation causes nuclear exclusion of Smads remains unknown. Agonists, including TGFb and BMP ligands, also induce phosphorylation of R-Smads at the linker region. While the antagonist-induced linker phosphorylation of Smads is cytoplasmic and dependent on MAP kinase, the one induced by agonists is mediated by cyclin-dependent kinases (CDKs), CDK8 and CDK9, in the nucleus, which are the components of transcriptional mediator and elongation complexes (Alarcon et al., 2009). Agonist-induced linker phosphorylation (ALP) of Smads occurs during or just prior to the association of Smads with transcriptional complexes. This ALP promotes Smad-dependent transcription while simultaneously marking the Smad proteins for proteasome-mediated turnover. In the case of Smad2 and Smad3, Nedd4L, an E3 ubiquitin ligase, is responsible for their degradation elicited by ALP (Gao et al., 2009). Nedd4L recognizes via its WW domain TGFb-induced phosphoThr-ProTyr motif in the linker region, resulting in polyubiquitination and turnover of Smad2 and 3. Furthermore, SGK1, a member of the PKB/Akt kinase family, has been shown to phosphorylate Nedd4L, inhibiting its binding to CDK8/9-phosphorylated Smad2 and 3 (Gao et al., 2009). Depletion of Nedd4L enhances the accumulation of C-terminally phosphorylated Smad2/3 as well as the induction of gene expression in response to TGFb/activin signals in human cell line and mouse embryonic stem cells (mESCs). This augmented sensitivity to activin signal in Nedd4L-depleted mESCs results in the induction of definitive endoderm, anterior mesoderm, and axial mesoderm at the expense of posterior mesoderm and extraembryonic mesoderm. Thus, it has been suggested that Nedd4L functions to limit Smad signaling in the TGFb and activin/nodal pathways.
D. Erbin Erbin (ErbB2/Her2-interacting protein) is a member of the leucine-rich repeat (LRR) and PDZ domain (LAP)-containing protein family. It associates specifically with Smad3 and Smad2 via a novel Smad-interacting domain (SID), thereby preventing Smad2/Smad3 from forming a complex with Smad4 (Dai et al., 2007). Thus, overexpression of Erbin suppresses TGFb/activin/Smad2-dependent, but not BMP/Smad1-mediated, transcriptional responses in Xenopus embryos and cultured cells, whereas its
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knockdown mediated by shRNA enhances cell growth inhibition and target gene expression induced by TGFb. Further studies are needed to dissect the physiological roles of Erbin during early development and cell homeostasis.
E. Transmembrane prostate androgen-induced RNA The transmembrane protein TMEPAI (transmembrane prostate androgeninduced RNA) is a direct early target gene of TGFb signaling. Overexpression and knockdown of TMEPAI inhibits and augments the induction of target genes and Smad2/3 phosphorylation, respectively, in response to TGFb and activin signals (Watanabe et al., 2010), suggesting its negative roles in these signaling. TMEPAI interacts directly with Smad2 and Smad3 through its Smad interaction motif (SIM). It has been demonstrated that TMEPAI competes with SARA, which recruits R-Smads to the activated receptor complex, for binding to Smad2 and Smad3, thereby sequestering these R-Smads from type I receptor activation (Watanabe et al., 2010). Thus, it has been suggested that TMEPAI functions to control the duration and intensity of TGFb and activin signaling through a negative feedback loop.
VI. Negative Regulation of Gene Transcription The activated Smad complexes bind to specific DNA sequences called Smad-binding element (SBE) in the promoters or enhancers of target genes. The intrinsic affinity of Smad proteins for their cognate DNA sequences is relatively low, and Smad complexes require site-specific DNA-binding cofactors to efficiently bind to the promoters (Feng and Derynck, 2005; Schmierer and Hill, 2007). FoxH1 (also known as FAST1), a winged-helix transcription factor, and Mixer and Milk, the Mix family members of homeodomain proteins, are well-known examples of Smad2-interacting transcription factors. These two transcriptional factors have distinct DNAbinding specificities and expression patterns. Tissue- or cell type-specific regulation of transcription by Smads also depends on interaction with additional transcriptional coactivators and control of chromatin remodeling activity associated with histone acetyltransferases, p300 and CREB-binding protein (CBP) (Feng and Derynck, 2005; Schmierer and Hill, 2007). Thus, blockade of DNA binding of Smads or of the recruitment of transcriptional coactivators is employed to regulate negatively TGFb/activin-dependent transcriptional responses (Fig. 5.3). Ski and SnoN are important negative regulators of TGFb/activin as well as BMP signaling. Since Ski and SnoN do not have catalytic activities, they inhibit Smad activity through association with other cellular partners. Ski
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Figure 5.3 Negative nuclear regulation of activin signaling. While the C-terminally phosphorylated Smad2 and 3 are dephosphorylated by PPM1A in the nucleus, Smad4 is ubiquitinated by an E3 ubiquitin ligase, Ectodermin/TIF1g, dissociated from Smad2/3 and then exported from the nucleus, leading to termination of signaling. A cytoplasmic enzyme, FAM/USP9x deubiquitinates the exported Smad4, restoring its ability to form a complex with Smad2/3. p53 cooperates with Smad complex in driving gene transcription by binding to the cis-element adjacent to the Smad-binding site on a common promoter. This DNA binding of p53 is impeded by XFDL156. FGF/MAPK-stimulated CK1 phosphorylates p53, enabling it to interact with Smad complex. Ski and SnoN interfere with the formation of R-Smad and Smad4 complex or association of coactivators p300/CBP and R-Smads, and recruit corepressors such as HDAC to inhibit transcription. FoxH1, Mixer, and Milk facilitate the binding of Smad complex to its cognate target DNA, which is prevented by DRAP1, SRF, and EVi-1 cofactors. Evi-1 also recruits corepressors, CtBP and HDAC, to block histone acetylation (Ac). Activated c-Jun binds to a transcriptional corepressor, TGIF, promoting the association of R-Smad and TGIF and inhibiting the recruitment of p300/CBP to the Smad complex to attenuate gene responses. BEN represses transcriptions by constitutively binding to the activin-response element of a promoter with HDAC3, which is antagonized by the Smads–TFII-1 complex stimulated by activin signal.
and SnoN interact with Smad2, Smad3, and Smad4, and are recruited to the SBE sequences in TGFb-responsive promoters, repressing the ability of Smad complex to activate target genes (Luo, 2004). In addition, Ski and SnoN have been shown to inhibit TGFb/activin signaling by interfering with the formation of heteromeric R-Smad and Smad4 complex or with the binding of R-Smads to the transcriptional coactivator p300/CBP, and recruiting the transcriptional corepressors including histone deacetylases (HDACs), N-CoR, and mSin3A (Luo et al., 1999; Wu et al., 2002).
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Overexpression of Ski or SnoN blocks the growth inhibitory activity of TGFb signals. Thus, many human cancer cells express highly Ski or SnoN and are refractory to TGFb-induced growth arrest. DRAP1 has been shown to repress with Dr1 as a general factor basal transcription by RNA polymerase II. However, DRAP1 also acts as a transcriptional corepressor specific for activin/nodal signaling (Iratni et al., 2002). It interacts with FoxH1 via the DNA-binding region of the latter, preventing a FoxH1–Smad2–Smad4 complex (ARF) from binding to the activin-response element from the Xenopus Mix2 promoter. Loss of Drap1 in mice causes defective mesoderm formation during gastrulation, which is linked to the excess of nodal signaling activity. Serum response factor (SRF) is also an inhibitory transcriptional cofactor of activin/nodal signaling. Inhibition of SRF function enhances transcriptional responses induced by activin/nodal signals in mammalian cells and expands the mesoderm-inducing activity of this signaling further into the prospective ectodermal region of Xenopus embryo, which leads to abnormal mesoderm formation (Yun et al., 2007). Mechanically, SRF interferes with the formation of the Smad2–FAST1 complex, probably blocking the binding of activated Smad2 to its cognate sequence of target genes. Thus, SRF functions to limit the spread of morphogenetic signals by impeding the responses to activin/nodal signaling in vertebrate embryo. Evi-1, a zinc finger-containing transcriptional factor, represses BMP-, activin-, and TGFb-elicited activation of reporter genes and endogenous Smad7 expression. It interacts with Smad1, 2, 3, and 4. Although Evi-1 has been shown to interrupt DNA binding of Smad complex in inhibiting TGFb signaling, it alternatively recruits the corepressor CtBP and its associated HDACs to attenuate TGFb-induced histone acetylation in the negative control of Smad7 expression (Alliston et al., 2005). In this process, the Smad–CBP complex functions as a scaffolding protein to facilitate DNA binding of Evi-1. Activation of c-Jun N-terminal kinase ( JNK) cascade inhibits the Smad2–Smad4–FAST1 complex-dependent activation of a reporter gene driven by activin-response element (ARE). Upon ligand stimulation, the activated c-Jun binds to the nuclear transcriptional corepressor TG-interacting factor (TGIF) and promotes the association of Smad2 and TGIF, which could lead to inhibition of the assembly of Smad2 and the coactivator p300 and repression of Smad2 transcriptional activity (Pessah et al., 2001). Since JNK cascade is activated in response to TGFb stimulation, it has been proposed that cells would exhibit diverse patterns of transcriptional responses to TGFb, depending on the relative activation of Smads versus JNK cascade. BEN (also known as WSCR11) is a member of the TFII-I family of transcription factors. It constitutively binds to the ARE of the Goosecoid (Gsc) promoter even without ligand activation and possibly recruits
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HDAC3 to repress the transcription of Gsc (Ku et al., 2005). Upon TGFb/ activin stimulation, the transcription factor TFII-1 binds Smad2, is recruited to the ARE sequence, displaces BEN–HDAC3 complex, and induces its expression. Thus, it is suggested that the related transcription factors of TFII-I family have opposing effects on gene responses to TGFb/activin signaling. A tumor suppressor p53 has been shown to cooperate strongly with the activated Smad complex to positively regulate TGFb/activin-dependent transcriptions in mammalian cells as well as during mesoderm formation in Xenopus embryo (Cordenonsi et al., 2003). p53 interacts with Smad2 and 3 in vivo, and they are recruited to distinct cis-regulatory elements on a common promoter of a target gene, leading to synergistic activation of transcription. The interaction of p53 and Smad2 or Smad3 requires phosphorylation of p53 at Ser6 and Ser9 by casein kinase 1d (CK1d) or CK1e whose activity is induced by FGF–Ras–Erk–MAPK signaling (Cordenonsi et al., 2007). Thus, p53 functions as an integration node between receptor tyrosine kinase (RTK) signaling and TGFb/activin signaling. A recent study has shown that the Zn-finger protein XFDL156 inhibits p53 activity during Xenopus ectoderm specification (Sasai et al., 2008). It interacts with the C-terminal regulatory region of p53 and blocks the binding of p53 to its cognate target DNA site, thereby inhibiting p53-dependent gene induction and mesodermal differentiation. Thus, XFDL156 acts to limit the mesoderm-inducing activity of TGFb/activin signals to the mesoderm by controlling cell responsiveness to p53.
VII. Conclusions Despite much progress in our understanding of how activin signaling is negatively regulated to modulate its intensity and duration, some important questions remain unresolved. For example, signal-induced modifications of receptors and Smads, including phosphorylation and dephosphorylation of type I activin receptor and R-Smads, and ubiquitination and deubiquitination of Smad4, are key determinants of the strength and length of signaling. Although some regulatory proteins such as Smad7, PPM1A, TIF1g, and FAM have been shown to be responsible for these processes, it is not clear how the kinetics of these modifications is regulated and what the molecular basis for this control is. It has been shown that the duration of activin signaling depends on the time spent by the internalized activin–activin receptor complex in the endolysosomal pathway ( Jullien and Gurdon, 2005). The Smad7/Smurf2 complex abolishes this duration by targeting the receptors to the lysosome. Rap2 GTPase attenuates receptor turnover mediated by this complex and
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prolongs activin signaling (Choi et al., 2008), suggesting an antagonism between Rap2 and Smad7 in control of receptor trafficking and signal duration. However, the molecular mechanism underlying this antagonism awaits additional investigation. TGFb receptors internalize through two distinct routes, which have different outcomes. A clathrin-mediated endocytosis of receptors promotes activation of signaling, whereas a lipid-raft/caveolae-dependent endocytic pathway is associated with receptor degradation and termination of signaling (Di Guglielmo et al., 2003). Thus, the proper partitioning of receptors into two membrane compartments seems to be critical for fine-tuning the duration of TGFb signaling. Some regulators, including interleukin (IL)-6, ADAM12, heparan sulfate, and hyaluronan, have been shown to be implicated in partitioning TGFb receptors into lipid-raft or non-lipid-raft microdomain (Atfi et al., 2007; Chen et al., 2006; Ito et al., 2004; Zhang et al., 2005). However, it is not known how these regulators affect the membrane distribution of TGFb receptors. It remains to be further examined whether activin receptors are also partitioned into different microdomains for distinct fates of trafficking, and these molecules also work for distribution of activin receptors. The Activin/Smad pathway is affected by other signaling pathways such as FGF/MAPK signaling. FGF/MAPK signals induce not only the linker phosphorylation of R-Smads to turn off activin signaling, but also CK1mediated phosphorylation of p53 to promote activin-dependent transcriptional responses in a cell type or context-dependent manner. The molecular mechanisms exerting these opposing effects will also be important subjects of future investigation. It is well established that activin functions as a morphogen to induce concentration-dependent distinct gene responses during development. Therefore, it will be interesting to examine the mechanisms by which various negative regulators of activin signaling contribute to its morphogenetic effects as well as their developmental relevance.
ACKNOWLEDGMENTS We apologize to those whose works are not cited due to limitation of space. We thank H. Y. Lee for help with preparation of figures. This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MEST) (No. 20090092829).
REFERENCES Adkins, H. B., Bianco, C., Schiffer, S. G., Rayhorn, P., Zafari, M., Cheung, A. E., Orozco, O., Olson, D., De Luca, A., Chen, L. L., et al. (2003). Antibody blockade of the Cripto CFC domain suppresses tumor cell growth in vivo. J. Clin. Invest. 112, 575–587.
Negative Regulation of Activin Signal Transduction
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Alarcon, C., Zaromytidou, A. I., Xi, Q., Gao, S., Yu, J., Fujisawa, S., Barlas, A., Miller, A. N., Manova-Todorova, K., Macias, M. J., et al. (2009). Nuclear CDKs drive Smad transcriptional activation and turnover in BMP and TGF-beta pathways. Cell 139, 757–769. Alliston, T., Ko, T. C., Cao, Y., Liang, Y. Y., Feng, X. H., Chang, C., and Derynck, R. (2005). Repression of bone morphogenetic protein and activin-inducible transcription by Evi-1. J. Biol. Chem. 280, 24227–24237. Atfi, A., Dumont, E., Colland, F., Bonnier, D., L’Helgoualc’h, A., Prunier, C., Ferrand, N., Clement, B., Wewer, U. M., and Theret, N. (2007). The disintegrin and metalloproteinase ADAM12 contributes to TGF-beta signaling through interaction with the type II receptor. J. Cell Biol. 178, 201–208. Batut, J., Schmierer, B., Cao, J., Raftery, L. A., Hill, C. S., and Howell, M. (2008). Two highly related regulatory subunits of PP2A exert opposite effects on TGF-beta/Activin/ Nodal signalling. Development 135, 2927–2937. Bianco, C., Normanno, N., Salomon, D. S., and Ciardiello, F. (2004). Role of the cripto (EGF-CFC) family in embryogenesis and cancer. Growth Factors 22, 133–139. Bilezikjian, L. M., Blount, A. L., Donaldson, C. J., and Vale, W. W. (2006). Pituitary actions of ligands of the TGF-beta family: Activins and inhibins. Reproduction 132, 207–215. Chen, C. L., Huang, S. S., and Huang, J. S. (2006). Cellular heparan sulfate negatively modulates transforming growth factor-beta1 (TGF-beta1) responsiveness in epithelial cells. J. Biol. Chem. 281, 11506–11514. Cheng, S. K., Olale, F., Bennett, J. T., Brivanlou, A. H., and Schier, A. F. (2003). EGFCFC proteins are essential coreceptors for the TGF-beta signals Vg1 and GDF1. Genes Dev. 17, 31–36. Cheong, S. M., Kim, H., and Han, J. K. (2009). Identification of a novel negative regulator of activin/nodal signaling in mesendodermal formation of Xenopus embryos. J. Biol. Chem. 284, 17052–17060. Cheyette, B. N., Waxman, J. S., Miller, J. R., Takemaru, K., Sheldahl, L. C., Khlebtsova, N., Fox, E. P., Earnest, T., and Moon, R. T. (2002). Dapper, a Dishevelled-associated antagonist of beta-catenin and JNK signaling, is required for notochord formation. Dev. Cell 2, 449–461. Choi, S. C., Kim, G. H., Lee, S. J., Park, E., Yeo, C. Y., and Han, J. K. (2008). Regulation of activin/nodal signaling by Rap2-directed receptor trafficking. Dev. Cell 15, 49–61. Cordenonsi, M., Dupont, S., Maretto, S., Insinga, A., Imbriano, C., and Piccolo, S. (2003). Links between tumor suppressors: p53 is required for TGF-beta gene responses by cooperating with Smads. Cell 113, 301–314. Cordenonsi, M., Montagner, M., Adorno, M., Zacchigna, L., Martello, G., Mamidi, A., Soligo, S., Dupont, S., and Piccolo, S. (2007). Integration of TGF-beta and Ras/MAPK signaling through p53 phosphorylation. Science 315, 840–843. Dai, F., Chang, C., Lin, X., Dai, P., Mei, L., and Feng, X. H. (2007). Erbin inhibits transforming growth factor beta signaling through a novel Smad-interacting domain. Mol. Cell. Biol. 27, 6183–6194. Degen, W. G., Weterman, M. A., van Groningen, J. J., Cornelissen, I. M., Lemmers, J. P., Agterbos, M. A., Geurts van Kessel, A., Swart, G. W., and Bloemers, H. P. (1996). Expression of nma, a novel gene, inversely correlates with the metastatic potential of human melanoma cell lines and xenografts. Int. J. Cancer 65, 460–465. Di Guglielmo, G. M., Le Roy, C., Goodfellow, A. F., and Wrana, J. L. (2003). Distinct endocytic pathways regulate TGF-beta receptor signalling and turnover. Nat. Cell Biol. 5, 410–421. Dupont, S., Zacchigna, L., Cordenonsi, M., Soligo, S., Adorno, M., Rugge, M., and Piccolo, S. (2005). Germ-layer specification and control of cell growth by Ectodermin, a Smad4 ubiquitin ligase. Cell 121, 87–99.
100
Sun-Cheol Choi and Jin-Kwan Han
Dupont, S., Mamidi, A., Cordenonsi, M., Montagner, M., Zacchigna, L., Adorno, M., Martello, G., Stinchfield, M. J., Soligo, S., Morsut, L., et al. (2009). FAM/USP9x, a deubiquitinating enzyme essential for TGFbeta signaling, controls Smad4 monoubiquitination. Cell 136, 123–135. Escalona, R. M., Stenvers, K. L., Farnworth, P. G., Findlay, J. K., and Ooi, G. T. (2009). Reducing betaglycan expression by RNA interference (RNAi) attenuates inhibin bioactivity in LbetaT2 gonadotropes. Mol. Cell. Endocrinol. 307, 149–156. Feng, X. H., and Derynck, R. (2005). Specificity and versatility in TGF-beta signaling through Smads. Annu. Rev. Cell Dev. Biol. 21, 659–693. Funaba, M., Zimmerman, C. M., and Mathews, L. S. (2002). Modulation of Smad2mediated signaling by extracellular signal-regulated kinase. J. Biol. Chem. 277, 41361–41368. Gao, S., Alarcon, C., Sapkota, G., Rahman, S., Chen, P. Y., Goerner, N., Macias, M. J., Erdjument-Bromage, H., Tempst, P., and Massague, J. (2009). Ubiquitin ligase Nedd4L targets activated Smad2/3 to limit TGF-beta signaling. Mol. Cell 36, 457–468. Gray, P. C., Greenwald, J., Blount, A. L., Kunitake, K. S., Donaldson, C. J., Choe, S., and Vale, W. (2000). Identification of a binding site on the type II activin receptor for activin and inhibin. J. Biol. Chem. 275, 3206–3212. Gray, P. C., Bilezikjian, L. M., and Vale, W. (2002). Antagonism of activin by inhibin and inhibin receptors: A functional role for betaglycan. Mol. Cell. Endocrinol. 188, 254–260. Gray, P. C., Harrison, C. A., and Vale, W. (2003). Cripto forms a complex with activin and type II activin receptors and can block activin signaling. Proc. Natl. Acad. Sci. USA 100, 5193–5198. Grimm, O. H., and Gurdon, J. B. (2002). Nuclear exclusion of Smad2 is a mechanism leading to loss of competence. Nat. Cell Biol. 4, 519–522. Grotewold, L., Plum, M., Dildrop, R., Peters, T., and Ruther, U. (2001). Bambi is coexpressed with Bmp-4 during mouse embryogenesis. Mech. Dev. 100, 327–330. Hamada, H., Meno, C., Watanabe, D., and Saijoh, Y. (2002). Establishment of vertebrate left-right asymmetry. Nat. Rev. Genet. 3, 103–113. Harrison, C. A., Gray, P. C., Vale, W. W., and Robertson, D. M. (2005). Antagonists of activin signaling: Mechanisms and potential biological applications. Trends Endocrinol. Metab. 16, 73–78. Hashimoto, O., Nakamura, T., Shoji, H., Shimasaki, S., Hayashi, Y., and Sugino, H. (1997). A novel role of follistatin, an activin-binding protein, in the inhibition of activin action in rat pituitary cells. Endocytotic degradation of activin and its acceleration by follistatin associated with cell-surface heparan sulfate. J. Biol. Chem. 272, 13835–13842. He, W., Dorn, D. C., Erdjument-Bromage, H., Tempst, P., Moore, M. A., and Massague, J. (2006). Hematopoiesis controlled by distinct TIF1gamma and Smad4 branches of the TGFbeta pathway. Cell 125, 929–941. Iratni, R., Yan, Y. T., Chen, C., Ding, J., Zhang, Y., Price, S. M., Reinberg, D., and Shen, M. M. (2002). Inhibition of excess nodal signaling during mouse gastrulation by the transcriptional corepressor DRAP1. Science 298, 1996–1999. Ito, T., Williams, J. D., Fraser, D. J., and Phillips, A. O. (2004). Hyaluronan regulates transforming growth factor-beta1 receptor compartmentalization. J. Biol. Chem. 279, 25326–25332. Itoh, S., and ten Dijke, P. (2007). Negative regulation of TGF-beta receptor/Smad signal transduction. Curr. Opin. Cell Biol. 19, 176–184. Janssens, V., and Goris, J. (2001). Protein phosphatase 2A: A highly regulated family of serine/threonine phosphatases implicated in cell growth and signalling. Biochem. J. 353, 417–439. Jullien, J., and Gurdon, J. (2005). Morphogen gradient interpretation by a regulated trafficking step during ligand-receptor transduction. Genes Dev. 19, 2682–2694.
Negative Regulation of Activin Signal Transduction
101
Kavsak, P., Rasmussen, R. K., Causing, C. G., Bonni, S., Zhu, H., Thomsen, G. H., and Wrana, J. L. (2000). Smad7 binds to Smurf2 to form an E3 ubiquitin ligase that targets the TGF beta receptor for degradation. Mol. Cell 6, 1365–1375. Kelber, J. A., Shani, G., Booker, E. C., Vale, W. W., and Gray, P. C. (2008). Cripto is a noncompetitive activin antagonist that forms analogous signaling complexes with activin and nodal. J. Biol. Chem. 283, 4490–4500. Kelber, J. A., Panopoulos, A. D., Shani, G., Booker, E. C., Belmonte, J. C., Vale, W. W., and Gray, P. C. (2009). Blockade of Cripto binding to cell surface GRP78 inhibits oncogenic Cripto signaling via MAPK/PI3K and Smad2/3 pathways. Oncogene 28, 2324–2336. Keutmann, H. T., Schneyer, A. L., and Sidis, Y. (2004). The role of follistatin domains in follistatin biological action. Mol. Endocrinol. 18, 228–240. Kowanetz, M., Lonn, P., Vanlandewijck, M., Kowanetz, K., Heldin, C. H., and Moustakas, A. (2008). TGFbeta induces SIK to negatively regulate type I receptor kinase signaling. J. Cell Biol. 182, 655–662. Kretzschmar, M., Doody, J., Timokhina, I., and Massague, J. (1999). A mechanism of repression of TGFbeta/Smad signaling by oncogenic Ras. Genes Dev. 13, 804–816. Ku, M., Sokol, S. Y., Wu, J., Tussie-Luna, M. I., Roy, A. L., and Hata, A. (2005). Positive and negative regulation of the transforming growth factor beta/activin target gene goosecoid by the TFII-I family of transcription factors. Mol. Cell. Biol. 25, 7144–7157. Lebrun, J. J., and Vale, W. W. (1997). Activin and inhibin have antagonistic effects on ligand-dependent heteromerization of the type I and type II activin receptors and human erythroid differentiation. Mol. Cell. Biol. 17, 1682–1691. Lewis, K. A., Gray, P. C., Blount, A. L., MacConell, L. A., Wiater, E., Bilezikjian, L. M., and Vale, W. (2000). Betaglycan binds inhibin and can mediate functional antagonism of activin signalling. Nature 404, 411–414. Li, Z. D., Wu, Y., Bao, Y. L., Yu, C. L., Guan, L. L., Wang, Y. Z., Meng, X. Y., and Li, Y. X. (2009). Identification and characterization of human ARIP2 and its relation to breast cancer. Cytokine 46, 251–259. Lin, X., Duan, X., Liang, Y. Y., Su, Y., Wrighton, K. H., Long, J., Hu, M., Davis, C. M., Wang, J., Brunicardi, F. C., et al. (2006). PPM1A functions as a Smad phosphatase to terminate TGFbeta signaling. Cell 125, 915–928. Liu, Z. H., Tsuchida, K., Matsuzaki, T., Bao, Y. L., Kurisaki, A., and Sugino, H. (2006). Characterization of isoforms of activin receptor-interacting protein 2 that augment activin signaling. J. Endocrinol. 189, 409–421. Liu, H. Y., Chen, F. F., Ge, J. Y., Wang, Y. N., Zhang, C. H., Cui, X. L., Yu, F., Tai, G. X., and Liu, Z. H. (2009). Expression and localization of activin receptorinteracting protein 2 in mouse tissues. Gen. Comp. Endocrinol. 161, 276–282. Luo, K. (2004). Ski and SnoN: Negative regulators of TGF-beta signaling. Curr. Opin. Genet. Dev. 14, 65–70. Luo, K., Stroschein, S. L., Wang, W., Chen, D., Martens, E., Zhou, S., and Zhou, Q. (1999). The Ski oncoprotein interacts with the Smad proteins to repress TGFbeta signaling. Genes Dev. 13, 2196–2206. Martens, J. W., de Winter, J. P., Timmerman, M. A., McLuskey, A., van Schaik, R. H., Themmen, A. P., and de Jong, F. H. (1997). Inhibin interferes with activin signaling at the level of the activin receptor complex in Chinese hamster ovary cells. Endocrinology 138, 2928–2936. Mathews, L. S., and Vale, W. W. (1991). Expression cloning of an activin receptor, a predicted transmembrane serine kinase. Cell 65, 973–982. Matsuzaki, T., Hanai, S., Kishi, H., Liu, Z., Bao, Y., Kikuchi, A., Tsuchida, K., and Sugino, H. (2002). Regulation of endocytosis of activin type II receptors by a novel
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PDZ protein through Ral/Ral-binding protein 1-dependent pathway. J. Biol. Chem. 277, 19008–19018. Mitchell, H., Choudhury, A., Pagano, R. E., and Leof, E. B. (2004). Ligand-dependent and -independent transforming growth factor-beta receptor recycling regulated by clathrinmediated endocytosis and Rab11. Mol. Biol. Cell 15, 4166–4178. Miura, S., Takeshita, T., Asao, H., Kimura, Y., Murata, K., Sasaki, Y., Hanai, J. I., Beppu, H., Tsukazaki, T., Wrana, J. L., et al. (2000). Hgs (Hrs), a FYVE domain protein, is involved in Smad signaling through cooperation with SARA. Mol. Cell. Biol. 20, 9346–9355. Moustakas, A., and Heldin, C. H. (2009). The regulation of TGFbeta signal transduction. Development 136, 3699–3714. Onichtchouk, D., Chen, Y. G., Dosch, R., Gawantka, V., Delius, H., Massague, J., and Niehrs, C. (1999). Silencing of TGF-beta signalling by the pseudoreceptor BAMBI. Nature 401, 480–485. Pessah, M., Prunier, C., Marais, J., Ferrand, N., Mazars, A., Lallemand, F., Gauthier, J. M., and Atfi, A. (2001). c-Jun interacts with the corepressor TG-interacting factor (TGIF) to suppress Smad2 transcriptional activity. Proc. Natl. Acad. Sci. USA 98, 6198–6203. Saito, S., Sidis, Y., Mukherjee, A., Xia, Y., and Schneyer, A. (2005). Differential biosynthesis and intracellular transport of follistatin isoforms and follistatin-like-3. Endocrinology 146, 5052–5062. Sasai, N., Yakura, R., Kamiya, D., Nakazawa, Y., and Sasai, Y. (2008). Ectodermal factor restricts mesoderm differentiation by inhibiting p53. Cell 133, 878–890. Schier, A. F. (2003). Nodal signaling in vertebrate development. Annu. Rev. Cell Dev. Biol. 19, 589–621. Schmierer, B., and Hill, C. S. (2007). TGFbeta-SMAD signal transduction: Molecular specificity and functional flexibility. Nat. Rev. Mol. Cell Biol. 8, 970–982. Schneyer, A. L., Rzucidlo, D. A., Sluss, P. M., and Crowley, W. F., Jr. (1994). Characterization of unique binding kinetics of follistatin and activin or inhibin in serum. Endocrinology 135, 667–674. Schneyer, A., Tortoriello, D., Sidis, Y., Keutmann, H., Matsuzaki, T., and Holmes, W. (2001). Follistatin-related protein (FSRP): A new member of the follistatin gene family. Mol. Cell. Endocrinol. 180, 33–38. Sekiya, T., Adachi, S., Kohu, K., Yamada, T., Higuchi, O., Furukawa, Y., Nakamura, Y., Nakamura, T., Tashiro, K., Kuhara, S., et al. (2004a). Identification of BMP and activin membrane-bound inhibitor (BAMBI), an inhibitor of transforming growth factor-beta signaling, as a target of the beta-catenin pathway in colorectal tumor cells. J. Biol. Chem. 279, 6840–6846. Sekiya, T., Oda, T., Matsuura, K., and Akiyama, T. (2004b). Transcriptional regulation of the TGF-beta pseudoreceptor BAMBI by TGF-beta signaling. Biochem. Biophys. Res. Commun. 320, 680–684. Shi, W., Sun, C., He, B., Xiong, W., Shi, X., Yao, D., and Cao, X. (2004). GADD34-PP1c recruited by Smad7 dephosphorylates TGFbeta type I receptor. J. Cell Biol. 164, 291–300. Shoji, H., Tsuchida, K., Kishi, H., Yamakawa, N., Matsuzaki, T., Liu, Z., Nakamura, T., and Sugino, H. (2000). Identification and characterization of a PDZ protein that interacts with activin type II receptors. J. Biol. Chem. 275, 5485–5492. Sidis, Y., Schneyer, A. L., Sluss, P. M., Johnson, L. N., and Keutmann, H. T. (2001). Follistatin: Essential role for the N-terminal domain in activin binding and neutralization. J. Biol. Chem. 276, 17718–17726. Stamler, R., Keutmann, H. T., Sidis, Y., Kattamuri, C., Schneyer, A., and Thompson, T. B. (2008). The structure of FSTL3. activin A complex. Differential binding of N-terminal
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domains influences follistatin-type antagonist specificity. J. Biol. Chem. 283, 32831–32838. Strizzi, L., Bianco, C., Normanno, N., and Salomon, D. (2005). Cripto-1: A multifunctional modulator during embryogenesis and oncogenesis. Oncogene 24, 5731–5741. Su, Y., Zhang, L., Gao, X., Meng, F., Wen, J., Zhou, H., Meng, A., and Chen, Y. G. (2007). The evolutionally conserved activity of Dapper2 in antagonizing TGF-beta signaling. FASEB J. 21, 682–690. ten Dijke, P., and Hill, C. S. (2004). New insights into TGF-beta-Smad signalling. Trends Biochem. Sci. 29, 265–273. Thompson, T. B., Lerch, T. F., Cook, R. W., Woodruff, T. K., and Jardetzky, T. S. (2005). The structure of the follistatin:activin complex reveals antagonism of both type I and type II receptor binding. Dev. Cell 9, 535–543. Tortoriello, D. V., Sidis, Y., Holtzman, D. A., Holmes, W. E., and Schneyer, A. L. (2001). Human follistatin-related protein: A structural homologue of follistatin with nuclear localization. Endocrinology 142, 3426–3434. Tsang, M., Kim, R., de Caestecker, M. P., Kudoh, T., Roberts, A. B., and Dawid, I. B. (2000). Zebrafish nma is involved in TGFbeta family signaling. Genesis 28, 47–57. Tsuchida, K., Arai, K. Y., Kuramoto, Y., Yamakawa, N., Hasegawa, Y., and Sugino, H. (2000). Identification and characterization of a novel follistatin-like protein as a binding protein for the TGF-beta family. J. Biol. Chem. 275, 40788–40796. Tsuchida, K., Nakatani, M., Matsuzaki, T., Yamakawa, N., Liu, Z., Bao, Y., Arai, K. Y., Murakami, T., Takehara, Y., Kurisaki, A., et al. (2004). Novel factors in regulation of activin signaling. Mol. Cell. Endocrinol. 225, 1–8. Tsukazaki, T., Chiang, T. A., Davison, A. F., Attisano, L., and Wrana, J. L. (1998). SARA, a FYVE domain protein that recruits Smad2 to the TGFbeta receptor. Cell 95, 779–791. Valdimarsdottir, G., Goumans, M. J., Itoh, F., Itoh, S., Heldin, C. H., and ten Dijke, P. (2006). Smad7 and protein phosphatase 1alpha are critical determinants in the duration of TGF-beta/ALK1 signaling in endothelial cells. BMC Cell Biol. 7, 16. Watanabe, Y., Itoh, S., Goto, T., Ohnishi, E., Inamitsu, M., Itoh, F., Satoh, K., Wiercinska, E., Yang, W., Shi, L., et al. (2010). TMEPAI, a transmembrane TGFbeta-inducible protein, sequesters Smad proteins from active participation in TGF-beta signaling. Mol. Cell 37, 123–134. Welt, C., Sidis, Y., Keutmann, H., and Schneyer, A. (2002). Activins, inhibins, and follistatins: From endocrinology to signaling. A paradigm for the new millennium. Exp. Biol. Med. 227, 724–752. Wiater, E., Lewis, K. A., Donaldson, C., Vaughan, J., Bilezikjian, L., and Vale, W. (2009). Endogenous betaglycan is essential for high-potency inhibin antagonism in gonadotropes. Mol. Endocrinol. 23, 1033–1042. Wrighton, K. H., Lin, X., and Feng, X. H. (2008). Critical regulation of TGFbeta signaling by Hsp90. Proc. Natl. Acad. Sci. USA 105, 9244–9249. Wu, J. W., Krawitz, A. R., Chai, J., Li, W., Zhang, F., Luo, K., and Shi, Y. (2002). Structural mechanism of Smad4 recognition by the nuclear oncoprotein Ski: Insights on Ski-mediated repression of TGF-beta signaling. Cell 111, 357–367. Xia, Y., and Schneyer, A. L. (2009). The biology of activin: Recent advances in structure, regulation and function. J. Endocrinol. 202, 1–12. Yamaguchi, T., Kurisaki, A., Yamakawa, N., Minakuchi, K., and Sugino, H. (2006). FKBP12 functions as an adaptor of the Smad7-Smurf1 complex on activin type I receptor. J. Mol. Endocrinol. 36, 569–579. Yan, X., Lin, Z., Chen, F., Zhao, X., Chen, H., Ning, Y., and Chen, Y. G. (2009). Human BAMBI cooperates with Smad7 to inhibit transforming growth factor-beta signaling. J. Biol. Chem. 284, 30097–30104.
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Yeo, C., and Whitman, M. (2001). Nodal signals to Smads through Cripto-dependent and Cripto-independent mechanisms. Mol. Cell 7, 949–957. Yun, C. H., Choi, S. C., Park, E., Kim, S. J., Chung, A. S., Lee, H. K., Lee, H. J., and Han, J. K. (2007). Negative regulation of Activin/Nodal signaling by SRF during Xenopus gastrulation. Development 134, 769–777. Zhang, L., Zhou, H., Su, Y., Sun, Z., Zhang, H., Zhang, Y., Ning, Y., Chen, Y. G., and Meng, A. (2004). Zebrafish Dpr2 inhibits mesoderm induction by promoting degradation of nodal receptors. Science 306, 114–117. Zhang, X. L., Topley, N., Ito, T., and Phillips, A. (2005). Interleukin-6 regulation of transforming growth factor (TGF)-beta receptor compartmentalization and turnover enhances TGF-beta1 signaling. J. Biol. Chem. 280, 12239–12245. Zhang, S., Fei, T., Zhang, L., Zhang, R., Chen, F., Ning, Y., Han, Y., Feng, X. H., Meng, A., and Chen, Y. G. (2007). Smad7 antagonizes transforming growth factor beta signaling in the nucleus by interfering with functional Smad-DNA complex formation. Mol. Cell. Biol. 27, 4488–4499.
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Antagonism of Activin by Activin Chimeras Uwe Muenster,1 Radhika Korupolu, Ratindra Rastogi, Jessica Read, and Wolfgang H. Fischer Contents I. Introduction II. The Activin/BMP Receptor System A. Activins and their receptors B. Bone morphogenetic proteins and their receptors C. Structural studies III. Design of Chimeras IV. Assessment of Binding and Biological Properties of Chimeras A. ActRII binding B. Activin-like bioactivity of chimeras C. Antagonism D. BMP-like activity of ActA/BMP chimeras V. Summary, Conclusions, and Future Directions Acknowledgments References
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Abstract Activins are pluripotent hormones/growth factors that belong to the TGF-b superfamily of growth and differentiation factors (GDFs). They play a role in cell growth, differentiation and apoptosis, endocrine function, metabolism, wound repair, immune responses, homeostasis, mesoderm induction, bone growth, and many other biological processes. Activins and the related bone morphogenic proteins (BMPs) transduce their signal through two classes of single transmembrane receptors. The receptors possess intracellular serine/ threonine kinase domains. Signaling occurs when the constitutively active type II kinase domain phosphorylates the type I receptor, which upon activation, phosphorylates intracellular signaling molecules. To generate antagonistic Clayton Foundation Laboratories for Peptide Biology, The Salk Institute for Biological Studies, La Jolla, California, USA Present address: Bayer Healthcare, Global Drug Discovery, Pharmaceutical Development, Forschungszentrum Aprath, Wuppertal, Germany
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Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00006-8
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ligands, we generated chimeric molecules that disrupt the receptor interactions and thereby the phosphorylation events. The chimeras were designed based on available structural data to maintain high-affinity binding to type II receptors. The predicted type I receptor interaction region was replaced by residues present in inactive homologs or in related ligands with different type I receptor affinities. ß 2011 Elsevier Inc.
I. Introduction Activins belong to the TGF-b superfamily of growth factors which control a variety of physiological functions such as cell growth, differentiation and apoptosis, endocrine function, metabolism, wound repair, immune responses, homeostasis, mesoderm induction, bone growth, and many others (Massague, 1998; Risbridger et al., 2001; Schubert et al., 1990; Vale et al., 1990, 1986; Werner and Grose, 2003). The TGF-b family comprises at least 42 members in humans, (Shi and Massague, 2003) including activin, TGF-b, bone morphogenetic protein, growth and differentiation factor (GDF), and nodal proteins, which are all characterized by a distinct structural feature, namely, a cysteine knot scaffold (Vitt et al., 2001). Activins and most other members of the TGF-b superfamily are homoor heterodimers consisting of two subunits, which are linked by a single covalent disulfide bond. Several subtypes of activin subunits are known and termed A, B, C, D, and E. The dimeric activins are named after their subunits, that is, activin A for an activin consisting of two A subunits, or AB for one consisting of one A and one B subunit. Proteins that were characterized biochemically include activin A, activin B, activin AB, and activin C. It has not been unambiguously established whether proteins corresponding with activins D or E or other subunit combinations exist. The structures of several members of the TGF-b superfamily, including different TGF-b subtypes, activin A, and several bone morphogenic proteins (BMPs), have been determined by X-ray crystallography and NMR procedures. In addition, for some of these ligands, structures in their bound state with receptor domains are available. Activins bind to two different subtypes of receptors termed type I and type II. Both receptor molecules are single transmembrane proteins. The N-terminal domain is engaged in ligand binding, whereas the intracellular C-terminal domain exhibits serine/ threonine kinase activities. In order to signal, activins initially bind to type II receptors with high affinity. The type I receptor is recruited into the complex and the constitutive kinase activity of the type II receptor phosphorylates residues in the intracellular kinase domain of the type I receptor, thereby activating it. Intracellular signaling molecules known as Smads are then phosphorylated and translocated to the nucleus where they modulate
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transcriptional events. BMPs are able to interact with activin type II receptors, in addition to BMP type II receptors, with high affinity. The specificity of the biological response is determined by the type I receptor which in turn interacts with a specific set of Smad signaling molecules. Inspection of the available structural and mutagenesis data allowed us to hypothesize that specific regions of activin and BMP are involved in interaction with either the type I or type II receptor. The goal of our investigation was the generation of activin mutants that would bind their type II receptor with near wild-type affinity. By manipulating the residues involved in type I receptor interaction, we aimed at generating ligands that could act as antagonists or be able to interact with different type I receptors, thereby altering their biological activity.
II. The Activin/BMP Receptor System A. Activins and their receptors Activins were first isolated and characterized based on their ability to release the reproductive hormone FSH from anterior pituitary cells (Ling et al., 1986; Vale et al., 1986). Subsequently, activins were implicated in a variety of diverse biological activities. These include roles in nerve cell survival, wound healing, and cell growth and differentiation. The major forms of activin are dimers of two closely related peptide chains, ba and bb, that form the active disulfide-linked proteins, that is, activin A, activin AB, and activin B. Other forms of activin chains, including the bc, bd, and be proteins are not widely expressed and their biological roles remain unclear. From radioactively labeled ligand cross-linking experiments, it was known that activin bound to two receptor components. Based on their apparent molecular weights, these had been termed type I for the lower molecular weight and type II for the higher one. The first member of this class of receptors from mammals, the activin type II receptor, was identified by expression cloning based on its ability to bind activin (Mathews and Vale, 1991). The protein was determined to be a single transmembrane molecule with an N-terminal ligand-binding domain and an internal serine/threonine kinase domain. The type II receptor exists in two homologous forms, termed ActRII and ActRIIB. These are distinct gene products which function in a similar manner. Subsequently, several type I receptors were identified by homology cloning and termed activin-receptor like kinases (ALKs) (ten Dijke et al., 1993). Further studies showed that activin A predominantly interacts with ALK4 (also known as ActRIB), whereas activin B interacts with ALK7 (Bertolino et al., 2008; ten Dijke et al., 1994; Tsuchida et al., 1993). While activin binds its type II receptor with high affinity, it only interacts with the type I receptor after binding to the
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type II molecule. The type II receptor is a constitutively active kinase, whereas the type I receptor’s serine–threonine kinase activity is activated after being phosphorylated by the type II kinase. Phosphorylation of the type I receptor takes place in its unique GS domain near the intracellular juxtamembrane regions preceding the serine–threonine kinase domain. The activated type I kinase then in turn phosphorylates Smad signaling molecules, specifically Smad 2 and Smad 3 in the case of the canonical activin type I receptor ALK4. Smad 2/3, which are also termed receptor-activated Smads, then form a complex with Smad 4 and trigger transcriptional activation events after translocation to the nucleus. Activin signaling is highly regulated at multiple levels (Phillips, 2000). The extracellular binding proteins, follistatin and follistatin-related protein, bind the ligand with high affinity and render it unable to interact with its receptors (Tsuchida et al., 2000). At the cell surface, the pseudoreceptor “BMP and activin membrane bound inhibitor” (BAMBI, Onichtchouk et al., 1999) interacts with several of the TGF-b family type I receptors and inhibits the formation of active receptor signaling complexes. Inhibin, the natural activin antagonist, binds the activin type II receptors with high affinity once bound to betaglycan (Lewis et al., 2000), thus blocking the interaction with activin. Cripto, a member of the EGF-CFC (epidermal growth factor-Cripto/frl/cryptic) family, which attaches to the outer cell membrane and functions as a coreceptor for nodal signaling, also inhibits interaction of activin with its receptors (Gray et al., 2003). Further proteins that interact with activin type II receptor are ARIPs 1 and 2 (activin receptor-interacting proteins). These are PDZ (PSD-95/Disc-large/ZO1) protein–protein interaction domain-containing proteins, that can either inhibit or enhance activin signaling, depending on the isoform expressed (Liu et al., 2006; Shoji et al., 2000). Further regulation occurs at the intracellular level of the Smad proteins, where inhibitory Smads (Smad 6, 7) disrupt TGF-b protein signal transduction. In the case of activin, Smad 7 has been shown to block signaling and to protect hepatocytes from activin A-induced growth inhibition (Kanamaru et al., 2001). Other proteins, such as Smurf-type ubiquitin E3 ligases, Smad anchor for receptor activation (SARA) (Tsukazaki et al., 1998), and transcriptional coactivators and corepressors like CBP, p300, c-Ski, and SnoN, influence Smad activation, and thus influence activin-linked biologic effects (Chen et al., 2006).
B. Bone morphogenetic proteins and their receptors Bone morphogenetic proteins (BMPs) were first identified as molecules that induce bone and cartilage formation in rodents (Wozney et al., 1988). BMPs are a large family (more than 20 members) with complex and diverse roles both in development and adult life (Matzuk et al., 1996; Whitman, 1998;
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Wozney et al., 1988). BMPs can be further classified into several subgroups, including the BMP-2/4 group, the BMP-5/6/7/8 group (OP-1 [osteogenic protein-1] group), GDF-5/6/7 group, and the BMP-9/10 group based on their biological activity (Kawabata et al., 1998). Many proteins of the BMP-2/4, OP-1, and BMP-9/10 groups induce formation of bone and cartilage tissues in vivo, while GDF-5/6/7 induce cartilage and tendon-like tissues, but not bone-like tissue. BMP-2, BMP-6, and BMP-9 were shown to be most potent in the induction of alkaline phosphatase activity and osteocalcin expression in the murine pluripotent mesenchymal cell line C3H10T1/2, and to induce differentiation of mesenchymal progenitor cells into osteoblasts, among 14 different BMPs and GDFs (Cheng et al., 2003). BMPs, like activins, signal via type II and type I receptors (Nohe et al., 2002) and subsequently activate Smad proteins 1, 5, and 8, which in turn transmit signals into the cell nucleus (Heldin et al., 1997; Massague, 1998). Both types of receptors are needed to form a functional complex to initiate downstream signaling events. The oligomerization pattern of the BMP receptors was shown to be quite different from that of receptors used by other members of the TGF-b superfamily (Gilboa et al., 2000). The most prominent difference is the presence of BMP receptor oligomers, heteromeric complexes of BMPR-II/BMPR-IA or BMPR-II/BMPR-IB, at the cell surface prior to ligand binding. This finding suggests that the specific signals from such preformed complexes might be mediated by conformational changes within the subunits in the complex upon ligand binding. Interestingly, Nohe et al. (2002) demonstrated that the mode of BMP receptor oligomerization determines which BMP signaling pathway is activated. In their studies, a measurable level of BMP receptors is found as preformed hetero-oligomeric complexes prior to ligand binding. However, a majority of receptors were recruited into hetero-oligomeric complexes only after ligand addition, where BMP first bound to BMPR-II that then recruited a specific BMPR-I into the signaling complex. They reported that BMP binding to preformed receptor complexes activates the Smad signaling pathway, whereas BMP-induced recruitment of receptors activates a Smadindependent signaling pathway. In addition to the canonical pathway described above, several Smad-independent pathways have been shown to be activated, including the p38 MAPK pathway (Derynck and Zhang, 2003; Iwasaki et al., 1999). 1. BMP type I receptors BMPs can interact with two distinct type I receptors that are ALKs. The type I receptors of BMPs are termed BMPR-IA (ALK-3) and BMPR-IB (ALK-6). BMPR-IA is widely expressed in various types of cells, while expression of BMPR-IB shows a more restricted expression profile. Specificities of various BMPs binding to type I receptors are affected by type II receptors (Yu et al., 2005) and appear crucial for understanding the
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pleiotropic effects of BMP action. BMP-2 and BMP-4 bind to BMPR-IA and BMPR-IB (ten Dijke et al., 1994), whereas BMP-6 and BMP-7 bind strongly to ALK-2 and weakly to BMPR-IB. GDF-5 preferentially binds to BMPR-IB, but not to other receptors (Nishitoh et al., 1996). BMP-9 and BMP-10 bind to ALK-1 and ALK-2 (Brown et al., 2005; David et al., 2007; Scharpfenecker et al., 2007). BMP type I receptors are shared by certain other members of the TGF-b family (Goumans et al., 2003; Oh et al., 2000). For example, MIS (Mu¨llerian inhibiting substance) binds to ALK-2, BMPR-IA, and BMPR-IB in the presence of its specific type II receptor, MISR-II. 2. BMP type II receptors Although BMP type I receptors (ALK3, ALK6, and ALK2) are largely specific for BMP family, it is not true for type II receptors. BMPR-II binds only BMPs (Kawabata et al., 1998; Liu et al., 2006; Rosenzweig et al., 1995) but activin type II receptors can bind both BMPs and activins (Yamashita et al., 1995). For example, BMP-3 can bind to both ActRII and BMPR-II and serve as modulators of growth factor activity during embryonic development. BMPs can thus bind to three type II receptors, that is, BMPR-II, ActRII, and ActRIIB, in mammals and these are widely expressed in various tissues. BMPR-II is specific for BMPs, whereas ActRII and ActRIIB can bind activins, myostatin, and BMPs. These type II receptors appear to bind most BMP ligands and affect the binding preferences of BMPs to type I receptors (Yu et al., 2005). However, the majority of BMP signaling utilizes the BMPR-II receptor. BMPR-II has a unique, long Cterminal tail with 530 amino acids following the Ser/Thr kinase domain (Rosenzweig et al., 1995). The long form with the C-terminal tail is predominantly expressed in most types of cells, while the short form without the C-terminal tail may be expressed in certain types of cells, as the tail region is shown to be not essential for BMP signaling in Xenopous embryo (Ishikawa et al., 1995). 3. Coreceptors for BMPs Although type II and type I receptors are sufficient for transduction of intracellular signaling by BMPs, binding to receptors and signaling activity of certain ligands is regulated by coreceptors. For example, Glycosylphosphatidylinositol (GPI)-anchored proteins of the repulsive guidance molecule (RGM) family, including RGMa, b, and c, (RGMb and c are also known as DRAGON and hemojuvelin, respectively) act as coreceptors for BMP-2 and BMP-4, and enhance BMP signaling (Babitt et al., 2005, 2006; Samad et al., 2005). In mouse pulmonary artery smooth muscle cells, BMP-2/4 signaling requires BMPR-II, but not ActRII or ActRIIB. However, cells transfected with RGMa use both BMPR-II and ActRII for BMP-2/4 signaling, suggesting that RGMa facilitates the use of ActRII
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by BMP-2/4 (Xia et al., 2007). BMP-6 plays a key role in iron metabolism in hepatocytes. Upon BMP-6 stimulation in hepatocytes, hemojuvelin induces the expression of hepcidin, which in turn decreases iron absorption by the intestine and iron release from macrophages (Babitt et al., 2006), thus indicating that hemojuvelin functions as a signaling component in the BMP signaling pathway. Concurring this hypothesis, mutations in Hemojuvelin gene were identified in individuals with juvenile hemochromatosis (type IIA), which is characterized by accumulation of iron in various organs (Babitt et al., 2006) similar to the mutations in the Hepcidin gene in those humans. Another example is the transmembrane protein Endoglin, which is expressed in proliferating endothelial cells and other cell types, and binds to various ligands, including TGF-b 1/3, activin A, and BMP-2/7 (Barbara et al., 1999). Although its function in TGF-b family signaling has not been fully determined, ectopic expression of endoglin results in inhibition of TGF-b-induced responses, whereas it enhances BMP-7-induced responses (Scherner et al., 2007). Mutations in the human ENG gene (encoding endoglin) result in hereditary hemorrhagic telangiectasia (HHT1, also known as Osler–Weber–Rendu disease) similar to those in ALK1 (which induce HHT2), suggesting that they act in a common signaling pathway ( Johnson et al., 1996; McAllister et al., 1994).
C. Structural studies 1. Available complex structures Various crystal structures of ligands bound to their corresponding receptors’ extracellular binding domains (ECD) have been resolved. So far, activin bound to ActRIIBECD (Greenwald et al., 2004; Thompson et al., 2003), TGF-b3 bound to TbRIIECD (Hart et al., 2002) as well as a ternary complex of TGF-b3 bound to TbRIECD and TbRIIECD (Groppe et al., 2008), BMP-7 bound to ActRIIECD (Greenwald et al., 2003), BMP2 bound to BMPR-IAECD (Kirsch et al., 2000) as well as ternary complexes of BMP-2 bound to BMPR-IAECD and ActRIIECD (Allendorph et al., 2006) and BMP-2 bound to BMPR-IAECD and ActRIIBECD (Weber et al., 2007), and GDF-5 bound to BMPR-IBECD (Kotzsch et al., 2009) are available. 2. Architecture of complex structures Not surprisingly, resolved complex structures reveal that TGF-b protein family members share common features with respect to ligand and receptor structure, as well as with regard to the overall architecture of ligand– receptor complexes. In each monomer of the ligand dimer, two pairs of antiparallel b-strands stretch out from the cysteine core of the dimer to form short and long fingers. The characteristic curvature of these fingers creates concave and convex surfaces on the ligand, which enable interaction with
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respective type I and type II receptor-binding epitopes. At the base of the fingers, each monomer has an a-helix, which together with the prehelix loop and the inner concave surface of the fingers of the other monomer, form the “wrist” region (Greenwald et al., 2004; Thompson et al., 2003). Each ligand dimer binds two type II receptors and two type I receptors, which leads to respective ternary signaling complexes. 3. Receptor–receptor interaction Direct interaction between the receptor ECDs was not observed for BMP-2 and BMP type I and II receptors. In the BMP2:BMPR-IA complex (Kirsch et al., 2000), no two atoms of BMPR-IA-ECDs were closer than 35 A˚ to each other. Also, no contacts between ActRII-ECDs and BMPRIA-ECDs were observed when models of BMP7:ActRII and BMP2: BMPR-IA were combined to form the ternary complex of BMP7: ActRII:BMPR-IA (closest distance: ActRII–ActRII, 83 A˚; BMPR-IA– ˚ ). However, the gap between C-termini of ActRII-ECD BMPR-IA, 66 A and BMPR-IA-ECD was 27 A˚ only, leaving open the possibility of contact between transmembrane segments (Greenwald et al., 2003). A similar conclusion was drawn from the ternary complex structure of BMP2: BMPR-IA:ActRIIB (Weber et al., 2007). No direct contact between ActRIIB-ECDs and BMPR-IA-ECDs was observed, with the closest ˚ . Distances between C-termini were ActRIIB– proximity being 12 A ˚ ˚ ; and BMPR-IA-ActRIIB, ActRIIB, 85 A; BMPR-IA–BMPR-IA, 70 A ˚ 40 A, excluding contacts of all the four transmembrane helices; however, contact of BMPR-IA and ActRIIB transmembrane segments and thus an influence of transmembrane segment interaction on downstream signaling cannot be ruled out. Also in the activin:ActRII complex, no direct contact between the two receptor ECDs was observed, with the closest proximity ˚ (Thompson et al., 2003); however, whether or not receptor being 31 A interaction between ActRII/ActRIIB and ALK4 occurs, awaits structural data with the ALK4 extracellular domain. In contrast, TGF-b receptors type I and II ECDs are in close enough distance to make direct contact (Groppe et al., 2008; Hart et al., 2002). Based on this finding, Groppe et al. speculated that the markedly cooperative manner of the assembly resulting from the highly specific receptor–receptor interaction may contribute to a full biological response over a narrow concentration range, when compared to BMP signaling, which displays a wide dose–response range (Benchabane and Wrana, 2003; Groppe et al., 2008). 4. Ligand flexibility BMP-2 and BMP-7 appear to be rather rigid ligands, with only minor conformational changes upon binding to their respective receptors. It was shown that the global fold of BMP-2 is not affected by binding to both type I and type II receptors (Allendorph et al., 2006; Kirsch et al., 2000; Weber
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et al., 2007), with only small locally restricted changes in backbone and side chain conformations observed in the wrist and knuckle epitopes (Weber et al., 2007). Ligand binding to BMPR-IA caused an induced fit in the prehelix region of BMP-2 (Pro48-Asn56). Also, binding of the BMP2: BMPR-IA complex to the type II receptor ActRIIB caused a small but significant reorientation of BMPR-IA (Weber et al., 2007), which did not occur in the complex of BMP-2 bound to BMPR-IA and ActRII (Allendorph et al., 2006). Therefore, it has been speculated that the observed conformational change in BMPR-IA after type II receptor binding may be dependent on the nature of the type II receptor (Weber et al., 2007). In contrast, TGF-b3 and activin A exhibit a high degree of flexibility, with significant conformational differences observed between the bound and the unbound state (Greenwald et al., 2004; Harrington et al., 2006; Hart et al., 2002; Thompson et al., 2003). NMR relaxation methods revealed an inherent flexibility in the TGF-b3 molecule, with the open state predominating at pH 4 and below and the closed state at pH 5 and above (Bocharov et al., 2002; Mittl et al., 1996), which may contribute to a dynamic equilibrium between “open” and “closed” dimer architecture. The conformation also appears to depend on whether TGF-b3 is in complex with one pair of type II receptors (open conformation; Hart et al., 2002) or in the full ternary complex (closed conformation; Groppe et al., 2008). Similar explanations may be true for activins as well, since two different conformations of activin A have been described when bound to ActRIIB (Greenwald et al., 2004; Thompson et al., 2003). However, a ternary complex of activin bound to ActRII/ActrIIB and ALK4 has not been resolved yet. GDF-5 exhibits a somewhat intermediate flexibility when compared to the rigid BMPs, and the flexible activin A and TGF-b3 (Kotzsch et al., 2009). Considering the observed differences in ligand flexibility within the TGF-b superfamily, it was speculated that dimer mobility within ligand–receptor complexes may play a role in the quantitative aspects of signaling and SMAD protein phosphorylation (Kotzsch et al., 2009; Thompson et al., 2003), as it has been discussed earlier for the EPO receptor (Syed et al., 1998). 5. Binding epitopes The ActRII (Allendorph et al., 2006; Greenwald et al., 2003) and ActRIIB (Greenwald et al., 2004; Thompson et al., 2003; Weber et al., 2007) binding epitopes are located in the convex curvatures of the ligand fingers, which are called the knuckle epitopes. Interestingly, the ActRII binding site in the BMP7:ActRII complex (Greenwald et al., 2003) is slightly more shifted away from the fingertips, when compared to the ActRII binding site in the ternary complex BMP2:ActRII:BMPR-IA (Allendorph et al., 2006). Therefore, it was speculated that the observed shift of the ActRII binding site is dependent on the ligand and thus may represent a mechanism for generating ligand-specific receptor recognition (Weber et al., 2007).
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Ligand-specific receptor identification was also observed with TGF-b3, in which the TbRII binding site is located in the finger tips (Groppe et al., 2008; Hart et al., 2002), and therefore significantly differs from the BMP-2, BMP-7, and activin A knuckle epitopes that interact with ActRII and ActRIIB. The type I receptor-binding epitope was assigned to the respective ligands’ wrists in all complex structures involving type I receptors (Allendorph et al., 2006; Groppe et al., 2008; Kirsch et al., 2000; Kotzsch et al., 2009; Weber et al., 2007). A detailed list of the various ligands’ amino acids involved in type I and type II receptor-binding according to the resolved complex structures is depicted in the sequence alignment (Fig. 6.1). Additionally, numerous mutagenesis studies confirmed several of these amino acids to contribute to ligand–receptor interaction (Fig. 6.1). With respect to type I receptor binding, eight missense mutations in GDF5 are known that cause skeletal malformation (Kotzsch et al., 2009); among those are three that cluster in the type I receptor-binding epitope (R57L, deltaL56/S58T/H59L, and L60P).With the two latter, type I receptor binding was abrogated (GDF5 deltaL56/S58T/H59L) or greatly decreased (GDF5 L60P) in a binding assay using immobilized BMPR-IA and BMPRIB-ECDs, whereas GDF5 R57L exhibited enhanced and unchanged affinity for BMPR-IA and BMPR-IB, respectively (Kotzsch et al., 2009). Another study using immobilized BMPR-IA and BMPR-IB ectodomains showed that BMP-2 W28F, D53A, Y103A hardly changed binding affinity for BMPR-IA and BMPR-IB, when compared to wt BMP-2, whereas
Figure 6.1 Sequence alignment of activin A, activin C, BMP-2, BMP-7, GDF-5, and TGF-b3. The mature, human activin A, activin C, BMP-2, BMP-7, GDF-5, and TGF-b3 protein sequences were aligned using the Megalign program. Activin A wrist residues changed with activin C/BMP2/BMP7 are indicated by a black line above activin A; residues involved in type I receptor binding according to resolved crystal structures of respective ligand–receptor complexes are shaded green (Allendorph et al., 2006; Keller et al., 2004; Kirsch et al., 2000; Kotzsch et al., 2009), and those involved in type I receptor binding according to mutagenesis studies are indicated by blue circles (Harrison et al., 2004; Keller et al., 2004; Kotzsch et al., 2009); residues involved in type II receptor binding according to resolved crystal structures of respective ligand–receptor complexes are shaded red (Allendorph et al., 2006; Greenwald et al., 2003, 2004; Hart et al., 2002; Thompson et al., 2003), and those involved in type II receptor binding according to mutagenesis studies are indicated by red circles (Allendorph et al., 2006; Keller et al., 2004; Kirsch et al., 2000; Weber et al., 2007; Wuytens et al., 1999).
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BMP-2 D53P, S69R, and L51P mutants exhibited a reduced affinity for BMPR-IA and BMPR-IB, with the most drastic loss of binding observed for the L51P mutant. Additionally, on a functional level, BMP-2 L51P almost completely lost its ability to induce alkaline phosphatase activity in C2C12 cells (Keller et al., 2004). Looking at type II receptor binding, BMP-2 variants D34A, L90A, L100A, and S88A exhibited a significant reduction of binding affinity to their low-affinity receptors ActRII- and BMPR-II-ECDs, as well as a relevant decrease in their ability to induce alkaline phosphatase in C2C12 cells (Kirsch et al., 2000), when compared to wt BMP-2. The strongest effect was observed with BMP-2 D34A, which after binding to its high-affinity type I receptor, competed with wt BMP2 for type II receptor binding, and thus acted as a BMP-2 antagonist in the before-mentioned alkaline phosphatase assay using C2C12 cells (Kirsch et al., 2000). Furthermore, in an attempt to identify residues being responsible for high-affinity binding of activin A to ActRII/ActRIIB, BMP-2 residues were replaced by respective activin A amino acids that were suspected to contribute to ActRII/ActRIIB binding. Indeed, the BMP-2 variant BMP-2 L100K/N102D (BMP-2 numbering) exhibited a strong increase in binding affinity for ActRIIB and to a lesser extent for ActRII, highlighting the relevance of these residues for ligand affinity for ActRII/ ActRIIB (Weber et al., 2007). Earlier, BMP-2 variants S85R, E109R, and L100K were shown to increase receptor affinity for ActRII when compared to wt BMP-2 (Allendorph et al., 2006), and the K102E activin A mutant was shown to lose its ability to bind ActRII and ActRIIB, and turned out to be inactive in a functional assay (Wuytens et al., 1999), underlining the importance of K102 in activin A for high-affinity binding to ActRII/ ActRIIB.
III. Design of Chimeras Activin mutants retaining high affinity for the type II receptor ActRII but losing the ability to signal in an activin-like manner (e.g., by disruption of the type I receptor binding site) would be potential activin antagonists, and might reveal a general principle for the generation of desired antagonists of TGF-b superfamily proteins. Since numerous BMP-2 residues involved in ALK3 binding are located in the BMP-2 wrist (Allendorph et al., 2006; Keller et al., 2004; Kirsch et al., 2000), it was suspected that this region significantly contributes to type I receptor binding of activin as well. However, with respect to a recent finding showing that the introduction of single point mutations in the activin wrist epitope does not have any significant effect on activin activity (Harrison et al., 2004), it was decided to generate activin chimeras with more severe mutations in the wrist.
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The biologically less active activin C only shares 11 out of 32 amino acids in the wrist with activin A, which might contribute to its low activity and also to a lower affinity for ALK4 when compared to activin A. Therefore, four mutants were constructed in which each of the eight residues are changed from activin A to activin C (A/C 46–53, A/C 54–61, A/C 62–69, and A/C 70–78), and one chimera with the entire wrist changed from activin A to activin C (A/C 46–78). Furthermore, structural data and mutagenesis studies described above not only proved the wrist epitope to be involved in type I receptor binding, but also led to the assumption that the wrist epitope determines type I receptor specificity of the respective ligands. Following this hypothesis, activin A/BMP chimeras were constructed by exchanging the wrist region (residues 45–79 of activin A) to corresponding BMP-2 or BMP-7 residues. Subsequently, it was tested whether or not created chimeras are able to recruit BMP type I receptors.
IV. Assessment of Binding and Biological Properties of Chimeras A. ActRII binding Based on the available data from previous crystal structure and mutagenesis studies, which showed that the ligands’ type II receptor binding interfaces are located in the ligands’ fingers (Greenwald et al., 2003; Kirsch et al., 2000; Thompson et al., 2003; Wuytens et al., 1999), the ActA/C and ActA/BMP chimeras were predicted to retain ActRII binding affinity, because both had been manipulated only in the wrist region. To prove this hypothesis, the chimeras’ ability to displace 125I-activin A from 293T cells transfected with mouse ActRII cDNA was assessed. Displacement curves revealed that EC50 values of wt-activin A for ActRII ranged between 90 and 200 pM, and EC50 values for ActA/C, ActA/BMP2, and ActA/BMP7 chimeras were 262–545 , 270 , and 120 pM, respectively, indicating that the chimeras indubitably retained their affinity for ActRII (Fig. 6.2).
B. Activin-like bioactivity of chimeras As the chimeras retained their type II binding as predicted, the activin-like bioactivity of the ActA/C and ActA/BMP chimeras was determined next. This was done by assessing their ability to activate an A3-lux reporter gene in HEK 293T cells, in which wt-activin A itself at a concentration of 20 nM led to a 32-fold induction of luciferase activity (EC50 200 pM). A/C 46–53, 54–61, 62–69, and 70–78 (EC50s ranging from 113–862 pM) showed activities comparable to wt-activin A (EC50 ranging from 70–320 pmol, (Muenster et al., 2005)); however, the maximum activity of A/C 54–61, A/C 62–69, and
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Figure 6.2 Competitive binding displacement curves. HEK293T cells were transfected with ActRII and subjected to competitive binding with unlabeled wt ActA, activin A/C chimera, or the different ActA/BMP chimeras as described (Korupolu et al., 2008; Muenster et al., 2005). Displacement curves are shown for activin A/C chimera (A), ActA/BMP2 chimera (B), and ActA/BMP7 chimera (C). A standard displacement curve generated in the presence of various concentrations of unlabeled activin A is also shown (A, B, and C). The value of each point on the graph is a mean S.D. of three measurements. Each experiment was repeated at least three times.
70–78 was somewhat reduced when compared to wt-activin A. Only chimeras in which the entire wrist was changed from activin A to corresponding residues of activin C (A/C 46–78), BMP-2 (ActA/BMP2), and BMP-7 (ActA/BMP7) were devoid of significant activin-like bioactivity in concentrations of up to 40 nM, suggesting that the chimeras lost their ability to signal in an activin-like manner (Fig. 6.3). Furthermore, the ActA/C chimeras retained their affinities for activin-binding protein, Follistatin (Muenster et al., 2005), which is in line with data published by Fischer et al. (2003),
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who generated a deletion mutant of activin A (activin des85–109) that does not bind follistatin, and thereby revealing that the C-terminus of activin A is crucial for follistatin binding.
C. Antagonism As the ActA/C 46–78 chimera and the ActA/BMP chimeras bind to ActRII and at the same time are devoid of activin-like bioactivity, they represent potential antagonists of proteins that signal via ActRII/ALK4 pathway like
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activins and myostatin (Rebbapragada et al., 2003). Therefore, we determined the ability of the chimeras to block activin A and myostatin-induced A3-lux reporter activity in HEK 293T cells. The chimeras indeed inhibited A3-lux reporter induction in a concentration-dependent manner (Fig. 6.4). IC50 ranges for the inhibition of luciferase activity induced by 100 pM activin A were 1– 8 nM (Fig. 6.4A), 1–10 nM (Fig. 6.4B), and 4 nM (Fig. 6.4C) for Act A/C 46–78, ActA/BMP2, and ActA/BMP7, respectively. Also, luciferase activity induced by 500 pM myostatin was reduced with IC50 ranges of 1–5 nM (Fig. 6.4A), 1–5 nM (Fig. 6.4B), and 2.4 nM (ranging from 1–8 nM) for Act A/C 46–78, ActA/BMP2, and ActA/BMP7, respectively. The ActA/C 46–78 chimera could not block the luciferase activity induced by 50 pM TGF-b (Muenster et al., 2005), which is in line with the fact that TGF-b acts via its own type II receptor, TbRII, followed by the recruitment of ALK5. In contrast, both ActA/BMP chimeras were able to inhibit TGF-b induced luciferase activity (data not shown), which may be explained by BMP-like activity of the ActA/BMP chimeras (via introduced BMP wrist epitopes), thus leading to activation of Smads 1, 5, and 8, which are known to have a negative effect on TGF-b bioactivity (Goumans et al., 2003). The antagonistic properties of the ActA/C 46–78 and ActA/BMP chimeras were further substantiated through their significant inhibitory effect on basal and activin-induced FSH release from the mouse gonadotrope cell line LbT2 as well as from rat anterior pituitary cells in primary cell culture (Korupolu et al., 2008; Muenster et al., 2005).
D. BMP-like activity of ActA/BMP chimeras Structural data and mutagenesis studies not only proved the wrist epitope to be involved in type I receptor binding (Allendorph et al., 2006; Keller et al., 2004; Kirsch et al., 2000; Fig. 6.1 sequence alignment), but also led to the assumption that the wrist epitope determines type I receptor specificity of the respective ligands. Following this hypothesis, it was tested whether or not the ActA/BMP2 and ActA/BMP7 chimeras are able to recruit BMP type I receptors and signal in a BMP-like manner. BMP-like activity of the ActA/BMP chimeras was assessed by characterizing their ability to activate BRE-Luc in HepG2 cells. HepG2 cells were used as they represent a relevant model (Duncan and Watt, 2001; Song et al., 1995) expressing both ActRII and BMP type I receptors and thus is a BMPresponsive system (Song et al., 1995). In contrast to wt-activin A, wt BMP-2 and wt BMP-7 stimulated BRE-Luc in HepG2, indicating that BRE-Luc activation is BMP-specific (Fig. 6.5A,B). Wt BMP-7 at a concentration of 40 nM led to a 23-fold BRE-Luc induction (EC50, 5 nM), whereas wt BMP-2 at a concentration of 40 nM induced BRE-Luc 12-fold (EC50, 4 nM). The ActA/BMP chimeras stimulated the BRE-Luc in HepG2 cells with EC50 values of 10 nM for ActA/BMP2 and 1 nM for ActA/BMP7
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Figure 6.4 Antagonism of activin A and myostatin by chimeras. HEK293T cells were transfected with A3 lux, b-galactosidase, and FAST2, and treated with increasing doses of activin A/C, ActA/BMP2, or ActA/BMP7 chimera in the presence of 100 pM wt activin A or 500 pM myostatin for 16–24 h. Cells were lysed and assayed for luciferase and b-galactosidase activities as described (Korupolu et al., 2008; Muenster et al., 2005). Dose–response curves activin A/C þ 100 pM wt activin A (open triangles) and activin A/C þ 500 pM myostatin (closed downward triangles) (A), ActA/BMP2 þ 100 pM wt activin A (open circle) and Act/ABMP2 þ 500 pM myostatin (open downward triangles) (B), and ActA/BMP7 þ 100 pM wt activin A (open square) and ActA/ BMP7 þ 500 pM myostatin (open diamond) (C) were generated using Prism software. Curves represent the luciferase activity normalized to b-galactosidase activity. Each point on the graph is a mean S.D. of three measurements.
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(Fig. 6.5A,B). These data indicated that the ActA/BMP chimeras indeed signal in a BMP-like manner. Further, ActA/BMP chimeras also exhibited BMP-like activity in the myoblastic cell line, C2C12. Under differentiating conditions, the ActA/ BMP chimeras provoked a significant shift in the differentiation of C2C12 cells toward osteoblast-like cells expressing alkaline phosphatase. Also, both chimeras caused a substantial increase in Phospho-Smad1 but not PhosphoSmad2 in C2C12 cells, clearly establishing that the chimeras, when compared to wt-activin A, have switched their binding affinities from the activin type I receptor ALK4 toward BMP type I receptors (Korupolu et al., 2008).
V. Summary, Conclusions, and Future Directions Signaling by ligands in the TGF-b superfamily is highly complex. Numerous ligands interact with a variety of receptors, which in turn activate diverse intracellular signaling molecules. The investigation of chimeric ligand molecules can shed a light on how specificity is achieved in such a system. We examined ligands of the activin/BMP subfamily. These ligands exhibit promiscuity toward the type II receptors but recruit and activate specific type I receptors. Using guidance from available crystal
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structures, we were able to identify a region of the ligand molecule that is prominently involved in the interaction with type I receptors. When exchanging residues in this region in activin A by ones present in the inactive activin C subtype, the ligand still bound its type II receptor with near wild-type affinity, yet was unable to interact with type I receptors in a way that would lead to transduction of signal. Antagonism is thus achieved by occupying type II receptors and rendering them unavailable for wildtype activin binding (Fig. 6.6). Extending this model of activin/BMP subfamily signal transduction, we predicted that exchanging residues in the region by corresponding ones from BMP-2 or BMP-7, recruitment and activation of BMP-specific type I receptors, and concurrent Smad activation would occur. As demonstrated (Fig. 6.5), the BMP-specific pathways are activated and phosphorylation of Smad 1 instead of Smad 2/3 occurs. Furthermore, as expected from the activin A/C chimera, the activin/BMP chimera acted as activin and myostatin antagonists as well. Altogether, data demonstrate that the wrist region of TGF-b protein family members significantly contributes to type I receptor specificity, and that manipulation of this region allows the construction of antagonists of the respective wild-type ligands.
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With respect to the implication of TGF-b protein family members in the pathogenesis of metabolic diseases, such as diabetes and obesity, muscle wasting conditions like muscular dystrophy, sarcopenia, and cachexia, fibrosis, bone diseases such as osteoporosis, and cancer (Tsuchida et al., 2009), antagonists of TGF-b proteins represent highly interesting therapeutic options. Thus far, promising attempts to block signaling of TGF-b proteins for therapeutic interventions include type I receptor kinase inhibitors (ALK4, 5, and 7; Ehata et al., 2007; Hjelmeland et al., 2004), soluble forms of extracellular domains of ActRIIA (Deal, 2009; Pearsall et al., 2008) and ActRIIB (Lee et al., 2005) both fused with IgG-Fc. Furthermore, overexpression of activin C (Gold et al., 2009) and ALK2 (Renlund et al., 2007) was demonstrated to antagonize activin effects in vitro. With the approach to create TGF-b protein antagonists presented in this publication, further work to characterize such antagonists needs to be done. It remains to be established whether changing the entire wrist between different members of the TGF-b superfamily would allow the construction of desired type II receptor antagonists. Additionally, further improvement of antagonistic properties might be achievable by increasing the affinity of the chimera for ActRII (Wuytens et al., 1999). Ultimately, in vivo experiments need to be performed in order to check whether the hypothesized biological effects of such antagonists will show.
ACKNOWLEDGMENTS We thank Drs. Wylie Vale, Ezra Wiater, Peter Gray, Craig Harrison, and Karsten Schmidt at the Salk Institute for discussions. We also thank Joan Vaughan and Cindy Donaldson for performing bioassays and for providing antibodies. This research was funded by a grant from NIH (HD135270) and by the Foundation for Medical Research.
REFERENCES Allendorph, G. P., Vale, W. W., and Choe, S. (2006). Structure of the ternary signaling complex of a TGF-beta superfamily member. Proc. Natl. Acad. Sci. USA 103, 7643–7648. Babitt, J. L., Zhang, Y., Samad, T. A., Xia, Y., Tang, J., Campagna, J. A., Schneyer, A. L., Woolf, C. J., and Lin, H. Y. (2005). Repulsive guidance molecule (RGMa), a DRAGON homologue, is a bone morphogenetic protein co-receptor. J. Biol. Chem. 280, 29820–29827. Babitt, J. L., Huang, F. W., Wrighting, D. M., Xia, Y., Sidis, Y., Samad, T. A., Campagna, J. A., Chung, R. T., Schneyer, A. L., Woolf, C. J., Andrews, N. C., and Lin, H. Y. (2006). Bone morphogenetic protein signaling by hemojuvelin regulates hepcidin expression. Nat. Genet. 38, 531–539. Barbara, N. P., Wrana, J. L., and Letarte, M. (1999). Endoglin is an accessory protein that interacts with the signaling receptor complex of multiple members of the transforming growth factor-beta superfamily. J. Biol. Chem. 274, 584–594.
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Benchabane, H., and Wrana, J. L. (2003). GATA- and Smad1-dependent enhancers in the Smad7 gene differentially interpret bone morphogenetic protein concentrations. Mol. Cell. Biol. 23, 6646–6661. Bertolino, P., Holmberg, R., Reissmann, E., Andersson, O., Berggren, P. O., and Ibanez, C. F. (2008). Activin B receptor ALK7 is a negative regulator of pancreatic beta-cell function. Proc. Natl. Acad. Sci. USA 105, 7246–7251. Bocharov, E. V., Korzhnev, D. M., Blommers, M. J., Arvinte, T., Orekhov, V. Y., Billeter, M., and Arseniev, A. S. (2002). Dynamics-modulated biological activity of transforming growth factor beta3. J. Biol. Chem. 277, 46273–46279. Brown, M. A., Zhao, Q., Baker, K. A., Naik, C., Chen, C., Pukac, L., Singh, M., Tsareva, T., Parice, Y., Mahoney, A., Roschke, V., Sanyal, I., et al. (2005). Crystal structure of BMP-9 and functional interactions with pro-region and receptors. J. Biol. Chem. 280, 25111–25118. Chen, Y. G., Wang, Q., Lin, S. L., Chang, C. D., Chuang, J., and Ying, S. Y. (2006). Activin signaling and its role in regulation of cell proliferation, apoptosis, and carcinogenesis. Exp. Biol. Med. (Maywood) 231, 534–544. Cheng, H., Jiang, W., Phillips, F. M., Haydon, R. C., Peng, Y., Zhou, L., Luu, H. H., An, N., Breyer, B., Vanichakarn, P., Szatkowski, J. P., Park, J. Y., et al. (2003). Osteogenic activity of the fourteen types of human bone morphogenetic proteins (BMPs). J. Bone Joint Surg. Am. 85-A, 1544–1552. David, L., Mallet, C., Mazerbourg, S., Feige, J. J., and Bailly, S. (2007). Identification of BMP9 and BMP10 as functional activators of the orphan activin receptor-like kinase 1 (ALK1) in endothelial cells. Blood 109, 1953–1961. Deal, C. (2009). Potential new drug targets for osteoporosis. Nat. Clin. Pract. Rheumatol. 5, 20–27. Derynck, R., and Zhang, Y. E. (2003). Smad-dependent and Smad-independent pathways in TGF-beta family signalling. Nature 425, 577–584. Duncan, S. A., and Watt, A. J. (2001). BMPs on the road to hepatogenesis. Genes Dev. 15, 1879–1884. Ehata, S., Hanyu, A., Fujime, M., Katsuno, Y., Fukunaga, E., Goto, K., Ishikawa, Y., Nomura, K., Yokoo, H., Shimizu, T., Ogata, E., Miyazono, K., et al. (2007). Ki26894, a novel transforming growth factor-beta type I receptor kinase inhibitor, inhibits in vitro invasion and in vivo bone metastasis of a human breast cancer cell line. Cancer Sci. 98, 127–133. Fischer, W. H., Park, M., Donaldson, C., Wiater, E., Vaughan, J., Bilezikjian, L. M., and Vale, W. (2003). Residues in the C-terminal region of activin A determine specificity for follistatin and type II receptor binding. J. Endocrinol. 176, 61–68. Gilboa, L., Nohe, A., Geissendorfer, T., Sebald, W., Henis, Y. I., and Knaus, P. (2000). Bone morphogenetic protein receptor complexes on the surface of live cells: A new oligomerization mode for serine/threonine kinase receptors. Mol. Biol. Cell 11, 1023–1035. Gold, E., Jetly, N., O’Bryan, M. K., Meachem, S., Srinivasan, D., Behuria, S., SanchezPartida, L. G., Woodruff, T., Hedwards, S., Wang, H., McDougall, H., Casey, V., et al. (2009). Activin C antagonizes activin A in vitro and overexpression leads to pathologies in vivo. Am. J. Pathol. 174, 184–195. Goumans, M. J., Valdimarsdottir, G., Itoh, S., Lebrin, F., Larsson, J., Mummery, C., Karlsson, S., and ten Dijke, P. (2003). Activin receptor-like kinase (ALK)1 is an antagonistic mediator of lateral TGFbeta/ALK5 signaling. Mol. Cell 12, 817–828. Gray, P. C., Harrison, C. A., and Vale, W. (2003). Cripto forms a complex with activin and type II activin receptors and can block activin signaling. Proc. Natl. Acad. Sci. USA 100, 5193–5198.
Antagonism of Activin by Activin Chimeras
125
Greenwald, J., Groppe, J., Gray, P., Wiater, E., Kwiatkowski, W., Vale, W., and Choe, S. (2003). The BMP7/ActRII extracellular domain complex provides new insights into the cooperative nature of receptor assembly. Mol. Cell 11, 605–617. Greenwald, J., Vega, M. E., Allendorph, G. P., Fischer, W. H., Vale, W., and Choe, S. (2004). A flexible activin explains the membrane-dependent cooperative assembly of TGF-beta family receptors. Mol. Cell 15, 485–489. Groppe, J., Hinck, C. S., Samavarchi-Tehrani, P., Zubieta, C., Schuermann, J. P., Taylor, A. B., Schwarz, P. M., Wrana, J. L., and Hinck, A. P. (2008). Cooperative assembly of TGF-beta superfamily signaling complexes is mediated by two disparate mechanisms and distinct modes of receptor binding. Mol. Cell 29, 157–168. Harrington, A. E., Morris-Triggs, S. A., Ruotolo, B. T., Robinson, C. V., Ohnuma, S., and Hyvonen, M. (2006). Structural basis for the inhibition of activin signalling by follistatin. EMBO J. 25, 1035–1045. Harrison, C. A., Gray, P. C., Fischer, W. H., Donaldson, C., Choe, S., and Vale, W. (2004). An activin mutant with disrupted ALK4 binding blocks signaling via type II receptors. J. Biol. Chem. 279, 28036–28044. Hart, P. J., Deep, S., Taylor, A. B., Shu, Z., Hinck, C. S., and Hinck, A. P. (2002). Crystal structure of the human TbetaR2 ectodomain–TGF-beta3 complex. Nat. Struct. Biol. 9, 203–208. Heldin, C. H., Miyazono, K., and ten Dijke, P. (1997). TGF-beta signalling from cell membrane to nucleus through SMAD proteins. Nature 390, 465–471. Hjelmeland, M. D., Hjelmeland, A. B., Sathornsumetee, S., Reese, E. D., Herbstreith, M. H., Laping, N. J., Friedman, H. S., Bigner, D. D., Wang, X. F., and Rich, J. N. (2004). SB-431542, a small molecule transforming growth factor-betareceptor antagonist, inhibits human glioma cell line proliferation and motility. Mol. Cancer Ther. 3, 737–745. Ishikawa, T., Yoshioka, H., Ohuchi, H., Noji, S., and Nohno, T. (1995). Truncated type II receptor for BMP-4 induces secondary axial structures in Xenopus embryos. Biochem. Biophys. Res. Commun. 216, 26–33. Iwasaki, S., Iguchi, M., Watanabe, K., Hoshino, R., Tsujimoto, M., and Kohno, M. (1999). Specific activation of the p38 mitogen-activated protein kinase signaling pathway and induction of neurite outgrowth in PC12 cells by bone morphogenetic protein-2. J. Biol. Chem. 274, 26503–26510. Johnson, D. W., Berg, J. N., Baldwin, M. A., Gallione, C. J., Marondel, I., Yoon, S. J., Stenzel, T. T., Speer, M., Pericak-Vance, M. A., Diamond, A., Guttmacher, A. E., Jackson, C. E., et al. (1996). Mutations in the activin receptor-like kinase 1 gene in hereditary haemorrhagic telangiectasia type 2. Nat. Genet. 13, 189–195. Kanamaru, C., Yasuda, H., Takeda, M., Ueda, N., Suzuki, J., Tsuchida, T., Mashima, H., Ohnishi, H., and Fujita, T. (2001). Smad7 is induced by norepinephrine and protects rat hepatocytes from activin A-induced growth inhibition. J. Biol. Chem. 276, 45636–45641. Kawabata, M., Imamura, T., and Miyazono, K. (1998). Signal transduction by bone morphogenetic proteins. Cytokine Growth Factor Rev. 9, 49–61. Keller, S., Nickel, J., Zhang, J. L., Sebald, W., and Mueller, T. D. (2004). Molecular recognition of BMP-2 and BMP receptor IA. Nat. Struct. Mol. Biol. 11, 481–488. Kirsch, T., Sebald, W., and Dreyer, M. K. (2000). Crystal structure of the BMP-2-BRIA ectodomain complex. Nat. Struct. Biol. 7, 492–496. Korupolu, R. V., Muenster, U., Read, J. D., Vale, W., and Fischer, W. H. (2008). Activin A/bone morphogenetic protein (BMP) chimeras exhibit BMP-like activity and antagonize activin and myostatin. J. Biol. Chem. 283, 3782–3790. Kotzsch, A., Nickel, J., Seher, A., Sebald, W., and Muller, T. D. (2009). Crystal structure analysis reveals a spring-loaded latch as molecular mechanism for GDF-5-type I receptor specificity. EMBO J. 28, 937–947.
126
Uwe Muenster et al.
Lee, S. J., Reed, L. A., Davies, M. V., Girgenrath, S., Goad, M. E., Tomkinson, K. N., Wright, J. F., Barker, C., Ehrmantraut, G., Holmstrom, J., Trowell, B., Gertz, B., et al. (2005). Regulation of muscle growth by multiple ligands signaling through activin type II receptors. Proc. Natl. Acad. Sci. USA 102, 18117–18122. Lewis, K. A., Gray, P. C., Blount, A. L., MacConell, L. A., Wiater, E., Bilezikjian, L. M., and Vale, W. (2000). Betaglycan binds inhibin and can mediate functional antagonism of activin signaling. Nature 404, 411–414. Ling, N., Ying, S.-Y., Ueno, N., Shimasaki, S., Esch, F., Hotta, M., and Guillemin, R. (1986). Pituitary FSH is released by a heterodimer of the beta-subunits from the two forms of inhibin. Nature 321, 779–782. Liu, Z. H., Tsuchida, K., Matsuzaki, T., Bao, Y. L., Kurisaki, A., and Sugino, H. (2006). Characterization of isoforms of activin receptor-interacting protein 2 that augment activin signaling. J. Endocrinol. 189, 409–421. Massague, J. (1998). TGF-beta signal transduction. Annu. Rev. Biochem. 67, 753–791. Mathews, L. S., and Vale, W. W. (1991). Expression cloning of an activin receptor, a predicted transmembrane serine kinase. Cell 65, 973–982. Matzuk, M. M., Kumar, T. R., Shou, W., Coerver, K. A., Lau, A. L., Behringer, R. R., and Finegold, M. J. (1996). Transgenic models to study the roles of inhibins and activins in reproduction, oncogenesis, and development. Recent Prog. Horm. Res. 51, 123–154, discussion 155-127. McAllister, K. A., Grogg, K. M., Johnson, D. W., Gallione, C. J., Baldwin, M. A., Jackson, C. E., Helmbold, E. A., Markel, D. S., McKinnon, W. C., Murrell, J., McCormick, M. K., Pericak-Vance, M. A., et al. (1994). Endoglin, a TGF-beta binding protein of endothelial cells, is the gene for hereditary haemorrhagic telangiectasia type 1. Nat. Genet. 8, 345–351. Mittl, P. R., Priestle, J. P., Cox, D. A., McMaster, G., Cerletti, N., and Grutter, M. G. (1996). The crystal structure of TGF-beta 3 and comparison to TGF-beta 2: Implications for receptor binding. Protein Sci. 5, 1261–1271. Muenster, U., Harrison, C. A., Donaldson, C., Vale, W., and Fischer, W. H. (2005). An activin-A/C chimera exhibits activin and myostatin antagonistic properties. J. Biol. Chem. 280, 36626–36632. Nishitoh, H., Ichijo, H., Kimura, M., Matsumoto, T., Makishima, F., Yamaguchi, A., Yamashita, H., Enomoto, S., and Miyazono, K. (1996). Identification of type I and type II serine/threonine kinase receptors for growth/differentiation factor-5. J. Biol. Chem. 271, 21345–21352. Nohe, A., Hassel, S., Ehrlich, M., Neubauer, F., Sebald, W., Henis, Y. I., and Knaus, P. (2002). The mode of bone morphogenetic protein (BMP) receptor oligomerization determines different BMP-2 signaling pathways. J. Biol. Chem. 277, 5330–5338. Oh, S. P., Seki, T., Goss, K. A., Imamura, T., Yi, Y., Donahoe, P. K., Li, L., Miyazono, K., ten Dijke, P., Kim, S., and Li, E. (2000). Activin receptor-like kinase 1 modulates transforming growth factor-beta 1 signaling in the regulation of angiogenesis. Proc. Natl. Acad. Sci. USA 97, 2626–2631. Onichtchouk, D., Chen, Y. G., Dosch, R., Gawantka, V., Delius, H., Massague, J., and Niehrs, C. (1999). Silencing of TGF-beta signalling by the pseudoreceptor BAMBI. Nature 401, 480–485. Pearsall, R. S., Canalis, E., Cornwall-Brady, M., Underwood, K. W., Haigis, B., Ucran, J., Kumar, R., Pobre, E., Grinberg, A., Werner, E. D., Glatt, V., Stadmeyer, L., et al. (2008). A soluble activin type IIA receptor induces bone formation and improves skeletal integrity. Proc. Natl. Acad. Sci. USA 105, 7082–7087. Phillips, D. J. (2000). Regulation of activin’s access to the cell: Why is mother nature such a control freak? Bioessays 22, 689–696.
Antagonism of Activin by Activin Chimeras
127
Rebbapragada, A., Benchabane, H., Wrana, J. L., Celeste, A. J., and Attisano, L. (2003). Myostatin signals through a transforming growth factor beta-like signaling pathway to block adipogenesis. Mol. Cell. Biol. 23, 7230–7242. Renlund, N., O’Neill, F. H., Zhang, L., Sidis, Y., and Teixeira, J. (2007). Activin receptorlike kinase-2 inhibits activin signaling by blocking the binding of activin to its type II receptor. J. Endocrinol. 195, 95–103. Risbridger, G. P., Schmitt, J. F., and Robertson, D. M. (2001). Activins and inhibins in endocrine and other tumors. Endocr. Rev. 22, 836–858. Rosenzweig, B. L., Imamura, T., Okadome, T., Cox, G. N., Yamashita, H., ten Dijke, P., Heldin, C. H., and Miyazono, K. (1995). Cloning and characterization of a human type II receptor for bone morphogenetic proteins. Proc. Natl. Acad. Sci. USA 92, 7632–7636. Samad, T. A., Rebbapragada, A., Bell, E., Zhang, Y., Sidis, Y., Jeong, S. J., Campagna, J. A., Perusini, S., Fabrizio, D. A., Schneyer, A. L., Lin, H. Y., Brivanlou, A. H., et al. (2005). DRAGON, a bone morphogenetic protein co-receptor. J. Biol. Chem. 280, 14122–14129. Scharpfenecker, M., van Dinther, M., Liu, Z., van Bezooijen, R. L., Zhao, Q., Pukac, L., Lowik, C. W., and ten Dijke, P. (2007). BMP-9 signals via ALK1 and inhibits bFGFinduced endothelial cell proliferation and VEGF-stimulated angiogenesis. J. Cell Sci. 120, 964–972. Scherner, O., Meurer, S. K., Tihaa, L., Gressner, A. M., and Weiskirchen, R. (2007). Endoglin differentially modulates antagonistic transforming growth factor-beta1 and BMP-7 signaling. J. Biol. Chem. 282, 13934–13943. Schubert, D., Kimura, H., LaCorbiere, M., Vaughan, J., Karr, D., and Fischer, W. H. (1990). Activin is a nerve cell survival molecule. Nature 344, 868–870. Shi, Y., and Massague, J. (2003). Mechanisms of TGF-beta signaling from cell membrane to the nucleus. Cell 113, 685–700. Shoji, H., Tsuchida, K., Kishi, H., Yamakawa, N., Matsuzaki, T., Liu, Z., Nakamura, T., and Sugino, H. (2000). Identification and characterization of a PDZ protein that interacts with activin type II receptors. J. Biol. Chem. 275, 5485–5492. Song, J. J., Celeste, A. J., Kong, F. M., Jirtle, R. L., Rosen, V., and Thies, R. S. (1995). Bone morphogenetic protein-9 binds to liver cells and stimulates proliferation. Endocrinology 136, 4293–4297. Syed, R. S., Reid, S. W., Li, C., Cheetham, J. C., Aoki, K. H., Liu, B., Zhan, H., Osslund, T. D., Chirino, A. J., Zhang, J., Finer-Moore, J., Elliott, S., et al. (1998). Efficiency of signalling through cytokine receptors depends critically on receptor orientation. Nature 395, 511–516. ten Dijke, P., Ichijo, H., Franzen, P., Schulz, P., Saras, J., Toyoshima, H., Heldin, C. H., and Miyazono, K. (1993). Activin receptor-like kinases: A novel subclass of cell-surface receptors with predicted serine/threonine kinase activity. Oncogene 8, 2879–2887. ten Dijke, P., Yamashita, H., Ichijo, H., Franzen, P., Laiho, M., Miyazono, K., and Heldin, C. H. (1994). Characterization of type I receptors for transforming growth factor–beta and activin. Science 264, 101–104. Thompson, T. B., Woodruff, T. K., and Jardetzky, T. S. (2003). Structures of an ActRIIB: Activin A complex reveal a novel binding mode for TGF-beta ligand:receptor interactions. EMBO J. 22, 1555–1566. Tsuchida, K., Mathews, L. S., and Vale, W. W. (1993). Cloning and characterization of a transmembrane serine kinase that acts as an activin type I reseptor. Proc. Natl. Acad. Sci. USA 90, 11242–11246. Tsuchida, K., Arai, K. Y., Kuramoto, Y., Yamakawa, N., Hasegawa, Y., and Sugino, H. (2000). Identification and characterization of a novel follistatin-like protein as a binding protein for the TGF-beta family. J. Biol. Chem. 275, 40788–40796.
128
Uwe Muenster et al.
Tsuchida, K., Nakatani, M., Hitachi, K., Uezumi, A., Sunada, Y., Ageta, H., and Inokuchi, K. (2009). Activin signaling as an emerging target for therapeutic interventions. Cell Commun. Signal 7, 15. Tsukazaki, T., Chiang, T. A., Davison, A. F., Attisano, L., and Wrana, J. (1998). SARA, a FYVE domain protein that recruits Smad2 to the TGF beta receptor. Cell 95, 779–791. Vale, W., Rivier, J., Vaughan, J., McClintock, R., Corrigan, A., Woo, W., Karr, D., and Spiess, J. (1986). Purification and characterization of an FSH releasing protein from porcine ovarian follicular fluid. Nature 321, 776–779. Vale, W., Hsueh, A., Rivier, C., and Yu, J. (1990). The inhibin/activin family of hormones and growth factors. In “Peptide Growth Factors and Their Receptors, Handbook of Experimental Pharmacology, Vol. 95/II,” (M. A. Sporn and A. B. Roberts, Eds.), pp. 211–248. Springer Verlag, Heidelberg. Vitt, U. A., Hsu, S. Y., and Hsueh, A. J. (2001). Evolution and classification of cystine knotcontaining hormones and related extracellular signaling molecules. Mol. Endocrinol. 15, 681–694. Weber, D., Kotzsch, A., Nickel, J., Harth, S., Seher, A., Mueller, U., Sebald, W., and Mueller, T. D. (2007). A silent H-bond can be mutationally activated for high-affinity interaction of BMP-2 and activin type IIB receptor. BMC Struct. Biol. 7, 6. Werner, S., and Grose, R. (2003). Regulation of wound healing by growth factors and cytokines. Physiol. Rev. 83, 835–870. Whitman, M. (1998). Smads and early developmental signaling by the TGFbeta superfamily. Genes Dev. 12, 2445–2462. Wozney, J. M., Rosen, V., Celeste, A. J., Mitsock, L. M., Whitters, M. J., Kriz, R. W., Hewick, R. M., and Wang, E. A. (1988). Novel regulators of bone formation: Molecular clones and activities. Science 242, 1528–1534. Wuytens, G., Verschueren, K., de Winter, J. P., Gajendran, N., Beek, L., Devos, K., Bosman, F., de Waele, P., Andries, M., van den Eijnden-van Raaij, A. J., Smith, J. C., and Huylebroeck, D. (1999). Identification of two amino acids in activin A that are important for biological activity and binding to the activin type II receptors. J. Biol. Chem. 274, 9821–9827. Xia, Y., Yu, P. B., Sidis, Y., Beppu, H., Bloch, K. D., Schneyer, A. L., and Lin, H. Y. (2007). Repulsive guidance molecule RGMa alters utilization of bone morphogenetic protein (BMP) type II receptors by BMP2 and BMP4. J. Biol. Chem. 282, 18129–18140. Yamashita, H., ten Dijke, P., Huylebroeck, D., Sampath, T. K., Andries, M., Smith, J. C., Heldin, C. H., and Miyazono, K. (1995). Osteogenic protein-1 binds to activin type II receptors and induces certain activin-like effects. J. Cell Biol. 130, 217–226. Yu, P. B., Beppu, H., Kawai, N., Li, E., and Bloch, K. D. (2005). Bone morphogenetic protein (BMP) type II receptor deletion reveals BMP ligand-specific gain of signaling in pulmonary artery smooth muscle cells. J. Biol. Chem. 280, 24443–24450.
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Activins and Cell Migration Hong-Yo Kang*,† and Chih-Rong Shyr‡,§ Contents I. Introduction II. Molecular Mechanism of Activin Signaling Regulated Cell Migration A. Smads-dependent cell migration B. Smads-independent cell migration III. The Role of Activins in the Regulation of Tumor Cell Migration and Metastasis A. Prostate cancer B. Breast cancer C. Colon cancer IV. The Role of Activins in the Modulation of Immune Cell Migration A. Mast cells B. Monocytes C. Dendritic cells V. Conclusion and Future Prospective Acknowledgments References
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Abstract Activins are the members of transforming growth factor b superfamily and act as secreted proteins; they were originally identified with a reproductive function, acting as endocrine-derived regulators of pituitary follicular stimulating hormone. In recent years, additional functions of activins have been discovered, including a regulatory role during crucial phases of growth, differentiation, and development such as wound healing, tissue repair, and regulation of branching morphogenesis. The functions of activins through activin receptors are pleiotrophic, while involving in the etiology and pathogenesis of a variety of diseases and being cell type-specific, they have been identified as important players in * Graduate Institute of Clinical Medical Sciences, Chang Gung University, College of Medicine, Kaohsiung, Taiwan Center for Menopause and Reproductive Research, Chang Gung Memorial Hospital-Kaohsiung Medical Center, Chang Gung University, College of Medicine, Kaohsiung, Taiwan { Department of Medical Laboratory Science and Biotechnology, China Medical University, Taichung, Taiwan } Sex Hormone Research Center, China Medical University Hospital, Taichung, Taiwan {
Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00007-X
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2011 Elsevier Inc. All rights reserved.
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cancer metastasis, immune responses, inflammation, and are most likely involved in cell migration. In this chapter, we highlight the current knowledge of activin signaling and discuss the potential physiological and pathological roles of activins acting on the migration of various cell types. ß 2011 Elsevier Inc.
I. Introduction Multicellular organisms require coordinated migration of cells to fulfill their needs in development and homeostasis. For instance, the three primitive germ layers are first formed during embryogenesis but later reestablish their relative positions during gastrulation; both physiological and pathological angiogenesis involve the movement of endothelial cells from preexisting vessels to new locations to form new vessels. On the other hand, tumor metastases can be interpreted as the result of invasion of the malignant cells to either adjacent or distant healthy tissues by means of lymphatic or hematologic spread, followed by extravasation and establishment of new tumor masses. The aforementioned processes are all intimately linked with cell migration, which is achieved via expressions of morphogens to persuade and guide movements of specific cell lineages down their genetically determined trajectories. Revealing mechanisms responsible for such precisely programmed migration may shed light on developing novel therapeutic drug targets for disease control. In contrast to bacterial chemotaxis, the mechanisms for eukaryotic cells to migrate involve multistage signal transduction, in which external chemotactic gradients are converted into intracellular signal gradients. The production of second messengers then begins and signaling cascades are activated, culminating in actin polymerization and the reorganization of cytoskeleton (Ribeiro et al., 2003). It is well established that extrinsic stimuli such as growth factors are critical in the process of cell migration, where stringent local cues such as target cell type, concentrations of soluble factors, and the surrounding microenvironment may all contribute (Ribeiro et al., 2003). Activins are members of the transforming growth factor beta (TGF-b) superfamily along with other multifunctional growth factors, comprising a subfamily of dimeric proteins consisting of two activin b units, which are linked by a disulfide bridge and contribute to cellular activity regulation (Xia and Schneyer, 2009). Activin A is a dimer of two activin bA subunits and has been identified to participate in a wide range of actions other than reproduction (Ball and Risbridger, 2001). Homodimer activin B (bB–bB) and heterodimer activin AB (bA–bB) of the subfamily also bind to their respective receptors and commence downstream signaling events (Robertson et al., 1992). Despite the fact that these different forms of activins
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are all bioactive, their nature vary, presenting distinct potencies and cellular functions, as demonstrated by in vitro assays with diverse cellular endpoints. The bc and bE subunit genes may encode proteins with antagonistic effects, leading to no downstream signaling whenever a homo- or heterodimer contains these subunits (Muenster et al., 2005), whereas each of the bA and bB subunits is capable of dimerizing with a structurally related but larger subunit to form inhibin A and B (de Kretser et al., 2000). The binding capacity of activin type II receptor by activins is lessened by binding of betaglycan by inhibins, which negatively regulate activities mediated by activins (de Kretser et al., 2000). Activins utilize a type I/type II receptor complex for signal transduction as other members of the TGF-b superfamily. Activin type II receptor (ACVR2 or ActRIIA) is a transmembrane protein harboring serine/threonine kinase activity for activin A (Mathews and Vale, 1991, 1993). A type II receptor of a different kind (ACVR2B or ActRIIB) has also been found (Harrison et al., 2004). Currently, seven type I receptors, activin receptorlike kinases 1–7 (ALK1–7) have been identified within the TGF-b family (Kang et al., 2009b). Type I receptors are no different from type II receptors in that they also possess the serine/threonine kinase activity; nevertheless, the uniqueness of type I receptors resides in the possession of the GS domain, which precedes the kinase domain and is close to the intracellular juxtamembrane regions. ALK4 is known as the activin type IB receptor (ACVR1B or ActRIB), while ALK7 is recognized as the activin type IC receptor (AVCR1C) (Graham and Peng, 2006). Type I receptors are recruited to the ligand/ActRII complex as soon as binding of activins to ActRIIA or ActRIIB occurs; this is followed by phosphorylation of GS domain by ActRII kinases (Graham and Peng, 2006). Subsequently, activated type I receptors phosphorylate activin/TGF-b-specific Smad, Smad2, and Smad3, which interact with the common mediator Smad4 to translocate into the nucleus for signal transduction initiation (Tsuchida et al., 2009). Although the DNA-binding property is intrinsic to Smads (Massague et al., 2005), various DNA-binding cofactors, including CBP/p300, TGIF, c-Ski, and Evi-1 (Kang et al., 2009b), associate with Smads in order to fully activate the target genes. There is an array of Smad-interacting transcription factors, ranging from members of the basic helix loop helix (bHLH) family, activator protein-1 (AP-1) family, and homeodomain protein family, to forkhead proteins and nuclear receptors (Derynck and Zhang, 2003). For aiding target gene regulation, additional transcriptional activators and repressors are also recruited to the Smad complexes once they are activated by activins/TGF-b signaling. These characteristics influence the specific patterns of transcription according to cell types and explain the level of complexity of activins/TGF-b signaling (Derynck and Zhang, 2003; Massague et al., 2005).
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II. Molecular Mechanism of Activin Signaling Regulated Cell Migration A. Smads-dependent cell migration Activin/TGF-b signaling impinges on the organization of cytoskeletal architecture in a rather complex manner. This action primarily involves the actin cytoskeleton and secondarily certain systems of intermediate filaments (Zhang et al., 2005). By targeting actin cytoskeleton, it probably aims at a minimum of two interconnected physiological manifestations: First, it facilitates cell motility, the prerequisites of which are the altered architectural arrangement and the remodeling of the extracellular matrix to which the cell adheres and migrates on. Second, the signaling alters the global architecture of the cell with further impact on its differentiation and proliferation. For those epithelial cells undergoing epithelial–mesenchymal transition (EMT), the alteration in cellular plasticity is particularly salient, which involves a modification in their differentiation program that increases the tendency for migration; this is essentially associated with the movements of tissues in the embryo and with tumor invasion and metastasis (Ball and Risbridger, 2001; Mercado-Pimentel and Runyan, 2007). During EMT, the intermediate filament system of cytokeratins exchanges to new cytokeratins and to a vimentin-based skeleton. These changes are functionally associated with induced cell mobility toward either a chemotactic gradient of TGF-b ligands or other member of TGF-b members. In the process of mesenchymal differentiation, Smad and the interacting transcription factors cooperate to provoke the expression of vimentin genes (Wu et al., 2007). As a result of vimentin synthesis, a new cytoskeleton of intermediate filaments, alpha-smooth muscle actin, and tropomyosins contributes to the assembly of new actomyosin networks that promote cell motility (Moustakas and Heldin, 2008). In addition, bone morphogenic protein (BMP) receptor complex recruitment and BMP-specific Smad signaling are activated and the new actomyosin networks are established upon myosin X synthesis in the filopodia of migrating cells (Moustakas and Heldin, 2008). Smad proteins are considered together with the regulation of actin dynamics and the modulation of Rho family GTPases, as exemplified by the interaction between Smad7 and ALK5 in prostate cancer cells, the former being inhibitory in nature and functions as a permissive factor for the activity of Cdc42 to be enhanced after its recruitment by ALK5 as an adaptor (Edlund et al., 2004). The role of Smad3 in EMT has been extensively reviewed recently (Xu et al., 2009; Zavadil and Bottinger, 2005). Consistent with these observations, kidney-derived primary tubular epithelial cells from Smad3 knockout mice are unable to enter EMT due to a failure of induction of vital regulators of transcription by TGF-b (Zavadil
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et al., 2004). Smad3-dependent EMT is blocked consistently by inhibitory Smad7 overexpression in vitro; this applies to pigment epithelium in the retina (Saika et al., 2004) and the epithelial cells of the mammary gland (Valcourt et al., 2005). EMT may also engage in the model of tumor progression, with Smad2 and/or Smad3 being implicated as the critical players, since the formation of metastasis and EMT mediating effects are attributed to the cooperation between Smad2 and H-ras (Oft et al., 2002). In addition, overexpression of Smad2 and Smad3 resulted in augmented EMT in a mammary epithelial model (Valcourt et al., 2005). These studies support that receptor-regulated Smads may have a central role in tumor progression and metastasis-associated EMT of TGF-b-dependent cells.
B. Smads-independent cell migration Intracellular pathways that are independent of Smad are also regulated, like their Smad-dependent counterparts, by the signaling through activin receptor complex upon ligand binding (Derynck and Zhang, 2003). p38 mitogen-activated protein kinase (MAPK), MAPK extracellular signal-regulated kinases (ERK) 1 and 2, and c-jun N-terminal kinase ( JNK) function specifically in their respective cell types and are downstream of activin signaling (Bao et al., 2005; Giehl et al., 2007); for instance, ERK1/2 activates the expression of tyrosine hydroxylase as a result of activin acting synergistically with basic fibroblast growth factor. Pituitary transcription factor Pit-1 is downregulated by activin via a pathway that is dependent on p38 MAPK, yet the presence of Smad is dispensable (de Guise et al., 2006). In the classical Wnt signaling pathway, the action of the coactivator ActRIB/Smad2 is independent of Smad4. Tcf4, b-catenin, and the coactivator p300 first come to close proximity with Smad2 after its activation, and then histone acetyltransferase activity that resides in p300 allows transcriptional enhancement of b-catenin/Tcf4 via physical interactions (Hirota et al., 2008). On top of Smads signaling, current evidences incline toward the idea that RhoGTPases and MAPK are also active in the activin-induced cell migration. The critical steps in activin-induced epithelial cell signal transductions occurring independent of Smad4 are ERK 1 and 2 and JNK activation. In activin-induced, Smad4-independent cell migration, attention has been given to the association of focal complexes with ERK and JNK, which may then exert significant effects along with the activated kinases. Unlike Rac and Cdc42, activation of RhoA can be achieved by activins and consequently leads to JNK and transcription factor c-Jun phosphorylation by MEKK1 (Zhang et al., 2005). p38 activity in keratinocytes from wildtype mice can be triggered by activin via a RhoA-independent pathway; however, such activity cannot be provoked in MEKK1-deficient mice. With their independence from Smad activation in mind, transcriptiondependent migration of keratinocytes due to activin stimulation still
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requires both p38 and MEKK1-mediated JNK activity (Zhang et al., 2005). The complexity of activin signaling is further emphasized by the regulation of focal contact turnover by recruited activated ERK and JNK and the reorganization of the dynamic cytoskeleton during activin-induced cell migration, which simultaneously widens the spectrum of activin-mediated, Smad4-independent cellular events participating in the course of action of cell migration.
III. The Role of Activins in the Regulation of Tumor Cell Migration and Metastasis The establishment of metastatic tumors is like tumorigenesis itself where a multistage process is necessary. The journey taken by the neoplastic cells from the primary tumor mass to the distant organs includes intravasation into, and survival within, the circulation, arrest, and extravasation into the secondary location; the maintenance of new colonies may be affected by the successful commencing and sustaining of growth and reinitiation of angiogenesis (Chambers et al., 2002). Therefore, only a minute amount of malignant cells leaving the primary tumor form metastatic mass eventually (Chambers et al., 2002). Genetic alterations is one of the fundamental requirements in the metastatic course, which pave the way for numerous changed cellular functions shown in both the malignant cells and the host tissues; these include the regulation of cell–cell adhesion, motility control, and interactions with the extracellular matrix ( Jacks and Weinberg, 2002). It is definitive that such genetic alterations may directly vary the expression patterns for certain genes; however, our knowledge to date with regard to the essence of tumor cell migration based on activin actions is still at its infancy.
A. Prostate cancer While activin A has been previously identified to inhibit the growth of prostate cancer cells and induces cell-cycle inhibitors such as p27 (Carey et al., 2004), the positive correlation between elevated serum activin A and prostate specific antigen (PSA) levels and increasing Gleason score in patients with bone-metastasizing prostate cancers is also well-documented (Incorvaia et al., 2007; Leto et al., 2006). The molecular mechanism underlying these two apparently paradoxical effects of activin A and how activin A influences the progression of prostate cancer with bone metastasis remains unclear. Based on our recent findings, we found that not only the expression of activin A is significantly increased in cancer biopsies with a bone metastatic propensity, but also activin A is a key factor promoting cancer cell
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migration to bone matrix produced by osteoblasts. We further discovered that activin A activates the androgen receptor (AR) function by modulating AR mRNA transcription and nuclear translocation through the Smad pathway, and that reduction of AR expression severely impaired the ability of activin A-treated cancer cells to migrate to bone matrix (Kang et al., 2009a). Hence, a model illustrating the role of activin A in prostate cancer metastasis is hypothesized (Fig. 7.1), in which two phases of downstream consequences of activin A’s action on prostate cancer cells may be characterized: (1) It increases PSA expression, cell migration, and adherence to bone matrix; these take place within a few hours of low dose activin A signaling and are counted toward short-term changes. (2) It triggers growth arrest for prostate cancer cells with altered cell morphology, and recruitment of cancer cells to bone lesions, with formation of osteoblastic matrix. These effects occur under prolonged treatment of activin A and are perceived as long-term changes. Among the short-term effects, AR activation through the Smad-dependent pathway is essential for cell motility enhancement. At the molecular level, activated AR has been shown to utilize the Src-FAKPI3 kinase-Cdc42/Rac1 cascade to mediate its cytoskeletal rearranging effects (Castoria et al., 2003). A number of cell surface proteins and proteins responsible for cell adhesion and migration, such as ezrin, integrin, and matrix metalloproteinase-2 (MMP-2), also demand AR for expression (Chuan et al., 2006; Nagakawa et al., 2004). Thus, following our evidences presented together with reports from others, it is conceivable to believe that prostate cancer cells’ metastatic predilection is linked with intrinsically determined gene expression, which requires strict synchronization of different signaling complexes and is susceptible to activin-mediated modulation through a pathway which may be put forth by the cooperation between AR and Smad proteins (Carey et al., 2004; Kang et al., 2001, 2002). On the other hand, ALK2-mediated phosphorylation of endoglin, a transmembrane glycoprotein that acts as a TGF-b co-receptor, has been reported to contribute to the regulation of prostate cancer cell migration through Smads-independent pathways (Craft et al., 2007). The present incomplete understanding of activin signaling on cell migration necessitates future work to dissect the involvement of activins/AR axis in the Cdc42/Rac1 polarity complex or other cascades related to ezrin, integrin, and MMP-2 to influence the motility of malignant prostate cells.
B. Breast cancer During the first week of lactation, activin A and its binding protein, follistatin, are found in human milk. In both primary and metastatic breast carcinoma, the gene expression of activin A subunit is more prominent
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Neuronendocrine cells 3 +
Blood vessel
1
Activin A 2
+ +
4
AR
5
Osteoblasts
Metastatic prostate cancer cells
6
New bone
Figure 7.1 Schematic illustration of activin A actions on the progression of prostate cancer. (1) Activin A is highly expressed in prostate cancers with metastatic capacity. (2) Activin A may be secreted as a paracrine factor to adjacent cells and blood vessels. (3) Activin A may induce the arrest of prostate cancer cells. (4) Activin A results in the expression of AR being upregulated by the cancer cells and promotes cell migration into the bone microenvironment. (5) Activin A is capable of mediating the adhesion of metastatic cancer cells at or near the cancer–bone interface. (6) Activin A is secreted by osteoblasts; this may result in further activation of metastatic prostate cancer cells. Blue numbers in the diagram illustrate how activin A works in our model, while gray numbers represent how activin A derived from the cancer and osteoblast cells may drive a vicious cycle of cancer metastasis.
compared to that of healthy tissues. When cell homogenates from breast cancer tissue are examined, the concentration of dimeric activin A is double the amount of that recorded from surrounding normal tissues. The precise role of activin A on breast cancer cells is so far waiting to be elucidated, and
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there is little understanding as to how it may entail these phenomena. Activins may suppress the growth of malignant cells, at least in the early stages of cancer progression. Both Cripto silencing and FLRG silencing techniques augment activin signaling and retrain the advancement of breast cancer cells (Adkins et al., 2003; Razanajaona et al., 2007). The responsiveness of breast cancer cells to activin A treatment can be divided into two classes depending on their expressions of estrogen receptors (ER). Previous studies revealed that the growths of cell lines that are deficient of ER expression were not inhibited by activin A; in fact, activin receptor expression in two of the cell lines proved to be low. In contrast, all of those that were positive for ER expression demonstrated activin A-dependent inhibition. (Kalkhoven et al., 1995). A conclusion drawn from the same study claims that the accentuated malignancy in ER-negative breast cancer cells relative to their ER-positive counterparts may be attributed to the poor responsiveness of activin signaling in the former (Kalkhoven et al., 1995). Nevertheless, it is too early a stage to assign activin with a cancer repression identity, and it is still uncertain in terms of the causative factors in either ER positive or negative cells of breast cancers taking the discrepancy in activin A actions into account. For breast and prostate cancer patients in later stages of disease, a remarkable elevation in serum activin concentration was observed and similarly, greater activin A level was evident in patients tested positive for bone metastasis, yet only the correlation between the number of bone metastases and activin A was considered significant (Leto et al., 2006). Interestingly, a prominent decline in activin A was indisputable during each of the first two days after tumor resection (Leto et al., 2006). Experimental inhibition of TGF-b reduced metastasis to various organs, including lung, liver, and bone (Ehata et al., 2007; Ogino et al., 2008; Talmadge, 2008), and the sensitivity of mammary epithelial cells to TGF-b stimulated migration was heightened and elicited by a crucial step which appears to be the phosphorylation of Smad1/5 (Giehl et al., 2007). Smad pathway is not the only signal transduction pathway identified that participates in the TGF-bdirected cell migration of epithelial breast carcinoma cell lines, JNK, RhoAGTPase activation, and the ERK 1 and 2 are also implicated (Giehl et al., 2007). Chemical inhibition targeting on ALK4 is a therapeutic strategy showing promising results, likewise similar effectiveness has also been observed in the model of in vivo bone metastasis (Ehata et al., 2007; Halder et al., 2005; Hjelmeland et al., 2004). Together these favor the idea that activins are associated with the pathogenesis of bone metastasis, and provides the rationale for using these cytokines for target therapy aiming primarily at preventing movement of malignant cells, hence combat bone metastasis. Another valuable aspect may be the use of activins as novel diagnostic markers for metastatic bone diseases.
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C. Colon cancer Microsatellite instability is one of the core genetic abnormalities found in colon cancers; 58% of 46 such cancer cell lines were identified to possess mutant ACVR2 ( Jung et al., 2007); the speculation of ACVR2 mutation being involved in colon carcinogenesis was further stressed by a separate study, showing an occurrence of mutations in the same gene, due to a frameshift in exon 10 in all 18 cases, with a proportion as great as 92% out of a total of 24 colon cancer cell lines and xenografts with high-frequency microsatellite instability (Plevova et al., 2004). In specimens for primary colon cancers, a comparable high rate of mutation was found paralleling the loss of ACVR2 protein expression in the majority of colon cancer cases ( Jung et al., 2006). In order to scrutinize the mechanisms and cellular effects of activin signaling in colon cancer, and to dissect possible new ACVR2 signaling transmission routes, ACVR2 function was rescued in two studies by cellular transfections, where a wild-type ACVR2 was introduced into tumor cell lines with high-frequency microsatellite instabilities that harbor the ACVR2 mutations. Under these circumstances, activin allowed ACVR2 protein to form a complex with ACVR1; subsequently, phosphorylated Smad2 was presented in the nucleus and activin-specific gene transcription was initiated. While reduced growth and S phase along with enhanced cellular migration were observed following activin treatment in the manipulated cells, small interfering RNA of ACVR2 reversed these effects (Deacu et al., 2004; Jung et al., 2007). Certain genes are thought to contribute to cell growth control and carcinogenesis, including the AP-1 complex genes JUND, JUN, and FOSB, and together with the members of the small GTPase signal transduction family, RHOB, ARHE, and ARHGDIA, their expression is amplified due to ACVR2 persuasion, as exhibited by the microarray-based differential expression analysis. Intriguingly, the overexpression of these genes coincides with TGF-b receptor 2 (TGFBR2) activation (Deacu et al., 2004). Thus, as one appreciates the analogous functional styles between activin and TGF-b signaling systems, it is not surprising for one to start assessing the possibility that activin signal may function as an alternative route through which the effectors of the TGF-b pathway are activated, with the phosphorylation of Smads being no exception. Regulation of activin-mediated responses may participate in the pathogenesis of high-frequency microsatellite instability colon cancers, considering its growth-restrictive and migration promotion effects akin to TGF-b in the colon cancer scenario. Together, the contribution of the activin signaling cascade to malignancy requires further evaluation to identify the synergies and differences to other members of the TGF-b superfamily.
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IV. The Role of Activins in the Modulation of Immune Cell Migration The migration of immune cells is critically important for the protective immune responses to tissues, which mainly involves the positioning of immunocompetent cells from bone marrow to the site of inflammation. Inflammation due to tissue damage or infection induces the release of cytokines and inflammatory chemoattractants from distressed stromal cells and sentinel cells, such as mast cells and macrophages, which then regulate immune cell migration by controlling the expressions of adhesion molecules for the trafficking of immune cells to the inflamed or damaged tissues (Stupack et al., 2000; Swart et al., 2005). The role of activins in the functioning of immune cells remained unclear. Recently, it is beginning to be revealed that activin A could act as a cytokine to regulate immune cell actions, including cell migration.
A. Mast cells Mast cells (MCs) are derived from multipotent hematopoietic progenitors in the bone marrow, involved in immediate hypersensitivity and chronic allergic reactions that can contribute to asthma, atopic dermatitis, and other allergic diseases (Okayama and Kawakami, 2006). MC migration is induced by various cytokines and chemokines to the sites of inflammation (Stupack et al., 2000), while activin A has autocrine effects on MCs. The induction of the activin bA gene in human MCs is stimulated by phorbol 12-myristate 13-acetate (PMA) and calcium ionophore (A23187; Cho et al., 2003), whereas the activation of the activin bA gene is achieved by the activation of JNK and p38 kinase through the calmodulin pathway in MCs (Funaba et al., 2003a). The expression of activin A may in turn modulates the function of MCs by the upregulating mouse mast cell protease-6 (mMCP-6), which is expressed in differentiated MCs (Funaba et al., 2005). With the evidences provided by the modulation of MC responses in experimental Smad3 depletion (Funaba et al., 2006), it is believed that Smad3-mediated signaling is essential for maximal cell growth in MCs. Activin A is present in murine bone marrow-derived, cultured mast cell progenitors (BMCMCs) expressing gene transcripts for molecules involved in activin signaling, suggesting that BMCMCs could be the target cells of activin A. Treatment of activin A inhibited cell growth of BMCMCs in a dose-dependent manner and caused morphological differentiation to upregulate the mRNA of mouse mast cell protease-1 (mMCP-1), a marker enzyme of mature mucosal MCs. Activin A showed significant activity in inducing the migration of BMCMCs; the optimal concentration for
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maximal migration was 10 pM, which was much lower than the concentrations required for inhibiting both cell growth and the activation of the mMCP-1 gene. Activin A secreted from activated immune cells recruits MC progenitors to sites of inflammation and that with increasing activin A concentrations, the progenitors differentiate into mature MCs. Thus, activin A may positively regulate the functions of MCs as effector cells of the immune system (Funaba et al., 2003b). The migratory response is probably mediated through its interaction with the TGF-b serine/threonine type I and II receptors in order to be expressed in the cells. This is supported by the evidence that TGF-b isoforms are highly potent chemotaxins for human MCs and can play an important role in the recruitment of MCs in inflammatory reactions (Olsson et al., 2000). Activin A enhances the transcription induced by Microphthalmia-associated transcription factor (MITF)-M and MITF-E, although MITF-mc blocked activin A-induced transcription of plasminogen activator inhibitor-1 (PAI-1), suggesting that discrete regulations of the plasminogen activator system occur in a cell type-specific manner (Murakami et al., 2006).
B. Monocytes Monocytes are equipped with phagocytic activity and the ability to differentiate into antigen-presenting cells to be involved in both innate and adaptive immune responses. Inflammatory cytokines play a role in regulating where and how do monocytes migrate. The expression of activin A in monocytes is highly regulated by inflammatory cytokines and glucocorticoids through a complex network of mechanisms (Abe et al., 2002; Dolter et al., 1998). Activin A expression is also stimulated by bacterial lipopolysaccharide (LPS) through protein kinase C-dependent transcriptional regulation (Eramaa et al., 1992), and the fact that activins increased the migrational activity of monocytes suggested a possible involvement of activins in regulating cell-mediated immune function (Petraglia et al., 1991). On the other hand, the proinflammatory cytokines such as IL-1, TGF-b, IFN-g, IL-8, and IL-10 also markedly enhance the expression of activin A mRNA in synoviocytes, suggesting their regulatory role in the control of activin A production in bone marrow stroma and monocytes (Yu et al., 1998). In addition, granulocyte–macrophage colony-stimulating factor (GM-CSF), glucocorticoids, or all-trans-retinoic acid were demonstrated to modulate the production of activin A by human monocytes ( Jaffe et al., 1995; Yu et al., 1996). Activin A inhibits the production of interleukin-1beta (IL-1b), a potent proinflammatory cytokine, and enhances the production of IL-1 receptor antagonist at the posttranscriptional level to act as an anti-inflammatory cytokine in inflammatory sites (Ohguchi et al., 1998). Activin A also induces TNF-a from monocytes, but in contrast, activin A has no effect on the production of TNF-b or IFN-g,
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both of which are known to be exclusively generated by T cells, indicating that activin A only plays a certain role in the physiological functions of normal human monocytes (Yamashita et al., 1993). Specifically, activin A induces the differentiation of human monocytes into Langerhans cells during inflammatory/autoimmune conditions (Musso et al., 2008).
C. Dendritic cells Dendritic cells (DCs) are derived from both myeloid and plasmacytoid DC (mDC and pDC) precursors. mDC precursors migrate to inflamed tissues in response to inflammatory chemokines and are then remobilized to regional lymph nodes, but pDC precursors transmigrate directly to regional lymph nodes via high endothelial venules (Yoneyama et al., 2005). DC acts as a source of activin A in vivo, monocyte-derived DC (Mo-DC) releases abundant levels of activin A during the maturation process induced by toll-like receptor (TLR) agonists, bacteria (Bartonella henselae, Salmonella thyphimurium), TNF, and CD40L (Scutera et al., 2008). Furthermore, activin A is also induced in monocyte-derived LC and in blood mDC by LPS and/or CD40L stimulation, but not in blood pDC following stimulation with influenza virus. Activin A production by DC is selectively downregulated by antiinflammatory molecules such as dexamethasone or IL-10. Neutralization of endogenous activin A using its inhibitor follistatin, or the addition of exogenous activin A during LPS treatment does not affect Mo-DC maturation marker expression, cytokine release, or allostimulatory function. However, Mo-DC matured with LPS in the presence of exogenous activin A displayed a higher FITC-dextran uptake, similar to that of immature DC (Scutera et al., 2008). Moreover, activin A promoted monocyte differentiation to DC and reversed the inhibitory effects of IL-6 on DC differentiation of monocytes. These findings demonstrate that activin A is released by different subsets of DC, and it is a cytokine that promotes DC generation, affecting the ability of mature DC to take up antigens (Ags; Scutera et al., 2008). Human Mo-DCs, the CD1c(þ) and CD123(þ) peripheral blood DC populations express both activin A and the type I and II activin receptors, rapidly secrete high levels of activin A after exposure to bacteria and specific TLR ligands, suggesting that activin A has potent autocrine effects on the capacity of human DCs to stimulate immune responses (Robson et al., 2008). Blocking autocrine activin A signaling in DCs using its antagonist, follistatin, enhanced DC cytokine (IL-6, IL-10, IL-12p70, and TNF-a) and chemokine (IL-8, IL-10, RANTES, and MCP-1) production during CD40L stimulation, but not TLR-4 ligation (Robson et al., 2008). Activin A induces the directional migration of immature myeloid dendritic cells (iDCs) through the activation of ALK4 and ActRIIA receptor chains by the selective and polarized release of two chemokines, namely, CXC chemokine ligands 12 and 14 through phosphatidylinositol 3-kinase gamma (Salogni et al., 2009).
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V. Conclusion and Future Prospective Although activins had been known as critical factors to stimulate follicle-stimulating hormone production from the anterior pituitary since the 1980s, an important role in cell migration processes for these proteins has only emerged more recently. In this chapter, we summarize the current knowledge of activin signaling and discuss the potential physiological and pathological roles of activins in various cell types during cell migration (Table 7.1). The significant elevation of the level of activin A in the serum of patients correlated with clinically evident prostate cancer metastases and PSA levels is particularly noteworthy, placing it as one of the earliest factors in a systemic cascade of cancer progression events. Nonetheless, its involvement in other tumor diseases like breast and colon cancers points to a pivotal function in all metastatic processes and also in associated pathological migration events. Further delineation of this group of protein complex during migration processes will allow their evaluation as potential diagnostic measures or therapeutic targets. Preliminary assessment of activins in various cellular migration pathologies may certainly be important, but a more thorough testing of their usefulness in monitoring and treating migration-related diseases such as cancer metastasis is warranted. It could be succinctly summarized as when, where, how, and why in terms of future delineation of the basic biology of these proteins in migration processes. The likely cellular timing for activin A-exerted effects is of prime importance particularly during migration processes, as we have argued that the altered cell behavior is likely to be solely accounted for by the rapid action of activin A, yet the more prolonged effects of activin A might also be required. Which cell types are influenced by these two phases and whether each phase takes place on distinct populations or has a more global impact is the subject of ongoing investigation. The mechanism by which activin A levels are elevated during migration processes is the second question, and it is a fundamental series of cellular and tissue events intimately linked with the timing of action of activin A. It is also of great interest to systematically identify activins’ target genes during migration processes, such as via microarray methods. This could help us to better understand the molecular basis of activins’ function and to design new strategies to combine antagonists with other drugs. For example, various strategies have been designed for the inhibition of activin signaling through receptors and soluble forms of the extracellular domains of activin receptors. Its natural binding protein, follistatin, and related ligand binding proteins, chemical kinase inhibitors for activin receptors, and siRNAs either for ligand or signaling molecules interfering with activin signaling, have also been suggested. Once promising proteins or chemicals targeting activin signaling are discovered, methods of
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Table 7.1 Summary of activin A action on various types of cells Cell types Cell name
Cancer cells
Activin A actions
Prostate Reduce cell growth; cancer cells promote cell migration and adherence to bone matrix; positively correlate with bone metastasis Breast cancer Reduce cell growth; cells positively correlate with bone metastasis Colon cancer Reduce cell growth and S cells phase along with enhanced cellular migration Immune Mast cells Inhibit BMCMCs’ cells growth; induce BMCMCs’ migration Monocytes Induce cell migration and differentiation; enhance cytokine production Dendritic cells Enhance maturation; promote differentiation; induce migration
References
Carey et al. (2004), Kang et al. (2009a)
Leto et al. (2006), Razanajaona et al. (2007) Deacu et al. (2004), Jung et al. (2007) Funaba et al. (2003b, 2006), Olsson et al. (2000) Ohguchi et al. (1998), Petraglia et al. (1991) Salogni et al. (2009), Scutera et al. (2008)
drug delivery are important issues for effective treatment. The final question, “why,” has produced some tantalizing hints, particularly from the findings that the dysregulation of activins may affect functions of gonads and adipose tissues. What this ultimately means in terms of the roles that the activins play in migration processes is yet to be deciphered, but the therapeutic interventions targeted to signaling through activin receptors may provide novel strategies for the development of effective treatments against a variety of diseases.
ACKNOWLEDGMENTS We apologize to the many researchers whose work could not be cited because of space limitations or was only cited indirectly by referring to reviews or more recent papers. This work was supported by grants CMRPD 87041, CMRPG 83021, and CMRPD 83038 from Chang Gung Memorial Hospital and NMRPD 140543 (NSC 94-2312-B-182-054) from the National Science Council to Dr. Hong-Yo Kang.
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REFERENCES Abe, M., Shintani, Y., Eto, Y., Harada, K., Kosaka, M., and Matsumoto, T. (2002). Potent induction of activin A secretion from monocytes and bone marrow stromal fibroblasts by cognate interaction with activated T cells. J. Leukoc. Biol. 72, 347–352. Adkins, H. B., Bianco, C., Schiffer, S. G., Rayhorn, P., Zafari, M., Cheung, A. E., Orozco, O., Olson, D., De Luca, A., Chen, L. L., Miatkowski, K., Benjamin, C., et al. (2003). Antibody blockade of the Cripto CFC domain suppresses tumor cell growth in vivo. J. Clin. Invest. 112, 575–587. Ball, E. M., and Risbridger, G. P. (2001). Activins as regulators of branching morphogenesis. Dev. Biol. 238, 1–12. Bao, Y. L., Tsuchida, K., Liu, B., Kurisaki, A., Matsuzaki, T., and Sugino, H. (2005). Synergistic activity of activin A and basic fibroblast growth factor on tyrosine hydroxylase expression through Smad3 and ERK1/ERK2 MAPK signaling pathways. J. Endocrinol. 184, 493–504. Carey, J. L., Sasur, L. M., Kawakubo, H., Gupta, V., Christian, B., Bailey, P. M., and Maheswaran, S. (2004). Mutually antagonistic effects of androgen and activin in the regulation of prostate cancer cell growth. Mol. Endocrinol. 18, 696–707. Castoria, G., Lombardi, M., Barone, M. V., Bilancio, A., Di Domenico, M., Bottero, D., Vitale, F., Migliaccio, A., and Auricchio, F. (2003). Androgen-stimulated DNA synthesis and cytoskeletal changes in fibroblasts by a nontranscriptional receptor action. J. Cell Biol. 161, 547–556. Chambers, A. F., Groom, A. C., and MacDonald, I. C. (2002). Dissemination and growth of cancer cells in metastatic sites. Nat. Rev. Cancer 2, 563–572. Cho, S. H., Yao, Z., Wang, S. W., Alban, R. F., Barbers, R. G., French, S. W., and Oh, C. K. (2003). Regulation of activin A expression in mast cells and asthma: its effect on the proliferation of human airway smooth muscle cells. J. Immunol. 170, 4045–4052. Chuan, Y. C., Pang, S. T., Cedazo-Minguez, A., Norstedt, G., Pousette, A., and FloresMorales, A. (2006). Androgen induction of prostate cancer cell invasion is mediated by ezrin. J. Biol. Chem. 281, 29938–29948. Craft, C. S., Romero, D., Vary, C. P., and Bergan, R. C. (2007). Endoglin inhibits prostate cancer motility via activation of the ALK2-Smad1 pathway. Oncogene 26, 7240–7250. de Guise, C., Lacerte, A., Rafiei, S., Reynaud, R., Roy, M., Brue, T., and Lebrun, J. J. (2006). Activin inhibits the human Pit-1 gene promoter through the p38 kinase pathway in a Smad-independent manner. Endocrinology 147, 4351–4362. de Kretser, D. M., Meinhardt, A., Meehan, T., Phillips, D. J., O’Bryan, M. K., and Loveland, K. A. (2000). The roles of inhibin and related peptides in gonadal function. Mol. Cell. Endocrinol. 161, 43–46. Deacu, E., Mori, Y., Sato, F., Yin, J., Olaru, A., Sterian, A., Xu, Y., Wang, S., Schulmann, K., Berki, A., Kan, T., Abraham, J. M., et al. (2004). Activin type II receptor restoration in ACVR2-deficient colon cancer cells induces transforming growth factorbeta response pathway genes. Cancer Res. 64, 7690–7696. Derynck, R., and Zhang, Y. E. (2003). Smad-dependent and Smad-independent pathways in TGF-beta family signalling. Nature 425, 577–584. Dolter, K. E., Palyash, J. C., Shao, L. E., and Yu, J. (1998). Analysis of activin A gene expression in human bone marrow stromal cells. J. Cell. Biochem. 70, 8–21. Edlund, S., Landstrom, M., Heldin, C. H., and Aspenstrom, P. (2004). Smad7 is required for TGF-beta-induced activation of the small GTPase Cdc42. J. Cell Sci. 117, 1835–1847. Ehata, S., Hanyu, A., Fujime, M., Katsuno, Y., Fukunaga, E., Goto, K., Ishikawa, Y., Nomura, K., Yokoo, H., Shimizu, T., Ogata, E., Miyazono, K., et al. (2007). Ki26894, a novel transforming growth factor-beta type I receptor kinase inhibitor, inhibits in vitro
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invasion and in vivo bone metastasis of a human breast cancer cell line. Cancer Sci. 98, 127–133. Eramaa, M., Hurme, M., Stenman, U. H., and Ritvos, O. (1992). Activin A/erythroid differentiation factor is induced during human monocyte activation. J. Exp. Med. 176, 1449–1452. Funaba, M., Ikeda, T., Murakami, M., Ogawa, K., and Abe, M. (2005). Up-regulation of mouse mast cell protease-6 gene by transforming growth factor-beta and activin in mast cell progenitors. Cell. Signal. 17, 121–128. Funaba, M., Ikeda, T., Ogawa, K., and Abe, M. (2003a). Calcium-regulated expression of activin A in RBL-2H3 mast cells. Cell. Signal. 15, 605–613. Funaba, M., Ikeda, T., Ogawa, K., Murakami, M., and Abe, M. (2003b). Role of activin A in murine mast cells: Modulation of cell growth, differentiation, and migration. J. Leukoc. Biol. 73, 793–801. Funaba, M., Nakaya, K., Ikeda, T., Murakami, M., Tsuchida, K., and Sugino, H. (2006). Requirement of Smad3 for mast cell growth. Cell. Immunol. 240, 47–52. Giehl, K., Imamichi, Y., and Menke, A. (2007). Smad4-independent TGF-beta signaling in tumor cell migration. Cells Tissues Organs 185, 123–130. Graham, H., and Peng, C. (2006). Activin receptor-like kinases: structure, function and clinical implications. Endocr. Metab. Immune Disord. Drug Targets 6, 45–58. Halder, S. K., Beauchamp, R. D., and Datta, P. K. (2005). A specific inhibitor of TGF-beta receptor kinase, SB-431542, as a potent antitumor agent for human cancers. Neoplasia 7, 509–521. Harrison, C. A., Wiater, E., Gray, P. C., Greenwald, J., Choe, S., and Vale, W. (2004). Modulation of activin and BMP signaling. Mol. Cell. Endocrinol. 225, 19–24. Hirota, M., Watanabe, K., Hamada, S., Sun, Y., Strizzi, L., Mancino, M., Nagaoka, T., Gonzales, M., Seno, M., Bianco, C., and Salomon, D. S. (2008). Smad2 functions as a co-activator of canonical Wnt/beta-catenin signaling pathway independent of Smad4 through histone acetyltransferase activity of p300. Cell. Signal. 20, 1632–1641. Hjelmeland, M. D., Hjelmeland, A. B., Sathornsumetee, S., Reese, E. D., Herbstreith, M. H., Laping, N. J., Friedman, H. S., Bigner, D. D., Wang, X. F., and Rich, J. N. (2004). SB-431542, a small molecule transforming growth factor-betareceptor antagonist, inhibits human glioma cell line proliferation and motility. Mol. Cancer Ther. 3, 737–745. Incorvaia, L., Badalamenti, G., Rini, G., Arcara, C., Fricano, S., Sferrazza, C., Di Trapani, D., Gebbia, N., and Leto, G. (2007). MMP-2, MMP-9 and activin A blood levels in patients with breast cancer or prostate cancer metastatic to the bone. Anticancer Res. 27, 1519–1525. Jacks, T., and Weinberg, R. A. (2002). Taking the study of cancer cell survival to a new dimension. Cell 111, 923–925. Jaffe, G. J., Roberts, W. L., Wong, H. L., Yurochko, A. D., and Cianciolo, G. J. (1995). Monocyte-induced cytokine expression in cultured human retinal pigment epithelial cells. Exp. Eye Res. 60, 533–543. Jung, B., Smith, E. J., Doctolero, R. T., Gervaz, P., Alonso, J. C., Miyai, K., Keku, T., Sandler, R. S., and Carethers, J. M. (2006). Influence of target gene mutations on survival, stage and histology in sporadic microsatellite unstable colon cancers. Int. J. Cancer 118, 2509–2513. Jung, B. H., Beck, S. E., Cabral, J., Chau, E., Cabrera, B. L., Fiorino, A., Smith, E. J., Bocanegra, M., and Carethers, J. M. (2007). Activin type 2 receptor restoration in MSIH colon cancer suppresses growth and enhances migration with activin. Gastroenterology 132, 633–644. Kalkhoven, E., Roelen, B. A., de Winter, J. P., Mummery, C. L., van den Eijnden-van Raaij, A. J., van der Saag, P. T., and van der Burg, B. (1995). Resistance to transforming
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growth factor beta and activin due to reduced receptor expression in human breast tumor cell lines. Cell Growth Differ. 6, 1151–1161. Kang, H. Y., Huang, H. Y., Hsieh, C. Y., Li, C. F., Shyr, C. R., Tsai, M. Y., Chang, C., Chuang, Y. C., and Huang, K. E. (2009a). Activin A enhances prostate cancer cell migration through activation of androgen receptor and is overexpressed in metastatic prostate cancer. J. Bone Miner. Res. 24, 1180–1193. Kang, H. Y., Huang, K. E., Chang, S. Y., Ma, W. L., Lin, W. J., and Chang, C. (2002). Differential modulation of androgen receptor-mediated transactivation by Smad3 and tumor suppressor Smad4. J. Biol. Chem. 277, 43749–43756. Kang, H. Y., Lin, H. K., Hu, Y. C., Yeh, S., Huang, K. E., and Chang, C. (2001). From transforming growth factor-beta signaling to androgen action: identification of Smad3 as an androgen receptor coregulator in prostate cancer cells. Proc. Natl. Acad. Sci. USA 98, 3018–3023. Kang, J. S., Liu, C., and Derynck, R. (2009b). New regulatory mechanisms of TGF-beta receptor function. Trends Cell Biol. 19, 385–394. Leto, G., Incorvaia, L., Badalamenti, G., Tumminello, F. M., Gebbia, N., Flandina, C., Crescimanno, M., and Rini, G. (2006). Activin A circulating levels in patients with bone metastasis from breast or prostate cancer. Clin. Exp. Metastasis 23, 117–122. Massague, J., Seoane, J., and Wotton, D. (2005). Smad transcription factors. Genes Dev. 19, 2783–2810. Mathews, L. S., and Vale, W. W. (1991). Expression cloning of an activin receptor, a predicted transmembrane serine kinase. Cell 65, 973–982. Mathews, L. S., and Vale, W. W. (1993). Molecular and functional characterization of activin receptors. Receptor 3, 173–181. Mercado-Pimentel, M. E., and Runyan, R. B. (2007). Multiple transforming growth factorbeta isoforms and receptors function during epithelial-mesenchymal cell transformation in the embryonic heart. Cells Tissues Organs 185, 146–156. Moustakas, A., and Heldin, C. H. (2008). Dynamic control of TGF-beta signaling and its links to the cytoskeleton. FEBS Lett. 582, 2051–2065. Muenster, U., Harrison, C. A., Donaldson, C., Vale, W., and Fischer, W. H. (2005). An activin-A/C chimera exhibits activin and myostatin antagonistic properties. J. Biol. Chem. 280, 36626–36632. Murakami, M., Ikeda, T., Saito, T., Ogawa, K., Nishino, Y., Nakaya, K., and Funaba, M. (2006). Transcriptional regulation of plasminogen activator inhibitor-1 by transforming growth factor-beta, activin A and microphthalmia-associated transcription factor. Cell. Signal. 18, 256–265. Musso, T., Scutera, S., Vermi, W., Daniele, R., Fornaro, M., Castagnoli, C., Alotto, D., Ravanini, M., Cambieri, I., Salogni, L., Elia, A. R., Giovarelli, M., et al. (2008). Activin A induces Langerhans cell differentiation in vitro and in human skin explants. PLoS ONE 3, e3271. Nagakawa, O., Akashi, T., Hayakawa, Y., Junicho, A., Koizumi, K., Fujiuchi, Y., Furuya, Y., Matsuda, T., Fuse, H., and Saiki, I. (2004). Differential expression of integrin subunits in DU-145/AR prostate cancer cells. Oncol. Rep. 12, 837–841. Oft, M., Akhurst, R. J., and Balmain, A. (2002). Metastasis is driven by sequential elevation of H-ras and Smad2 levels. Nat. Cell Biol. 4, 487–494. Ogino, H., Yano, S., Kakiuchi, S., Muguruma, H., Ikuta, K., Hanibuchi, M., Uehara, H., Tsuchida, K., Sugino, H., and Sone, S. (2008). Follistatin suppresses the production of experimental multiple-organ metastasis by small cell lung cancer cells in natural killer celldepleted SCID mice. Clin. Cancer Res. 14, 660–667. Ohguchi, M., Yamato, K., Ishihara, Y., Koide, M., Ueda, N., Okahashi, N., Noguchi, T., Kizaki, M., Ikeda, Y., Sugino, H., and Nisihara, T. (1998). Activin A regulates the
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production of mature interleukin-1beta and interleukin-1 receptor antagonist in human monocytic cells. J. Interferon Cytokine Res. 18, 491–498. Okayama, Y., and Kawakami, T. (2006). Development, migration, and survival of mast cells. Immunol. Res. 34, 97–115. Olsson, N., Piek, E., ten Dijke, P., and Nilsson, G. (2000). Human mast cell migration in response to members of the transforming growth factor-beta family. J. Leukoc. Biol. 67, 350–356. Petraglia, F., Sacerdote, P., Cossarizza, A., Angioni, S., Genazzani, A. D., Franceschi, C., Muscettola, M., and Grasso, G. (1991). Inhibin and activin modulate human monocyte chemotaxis and human lymphocyte interferon-gamma production. J. Clin. Endocrinol. Metab. 72, 496–502. Plevova, P., Krepelova, A., Papezova, M., Sedlakova, E., Curik, R., Foretova, L., Navratilova, M., Novotny, J., Zapletalova, J., Palas, J., Nieslanik, J., Horacek, J., et al. (2004). Immunohistochemical detection of the hMLH1 and hMSH2 proteins in hereditary non-polyposis colon cancer and sporadic colon cancer. Neoplasma 51, 275–284. Razanajaona, D., Joguet, S., Ay, A. S., Treilleux, I., Goddard-Leon, S., Bartholin, L., and Rimokh, R. (2007). Silencing of FLRG, an antagonist of activin, inhibits human breast tumor cell growth. Cancer Res. 67, 7223–7229. Ribeiro, C., Petit, V., and Affolter, M. (2003). Signaling systems, guided cell migration, and organogenesis: insights from genetic studies in Drosophila. Dev. Biol. 260, 1–8. Robertson, D. M., Foulds, L. M., Prisk, M., and Hedger, M. P. (1992). Inhibin/activin beta-subunit monomer: isolation and characterization. Endocrinology 130, 1680–1687. Robson, N. C., Phillips, D. J., McAlpine, T., Shin, A., Svobodova, S., Toy, T., Pillay, V., Kirkpatrick, N., Zanker, D., Wilson, K., Helling, I., Wei, H., et al. (2008). Activin-A: a novel dendritic cell-derived cytokine that potently attenuates CD40 ligand-specific cytokine and chemokine production. Blood 111, 2733–2743. Saika, S., Ikeda, K., Yamanaka, O., Sato, M., Muragaki, Y., Ohnishi, Y., Ooshima, A., Nakajima, Y., Namikawa, K., Kiyama, H., Flanders, K. C., and Roberts, A. B. (2004). Transient adenoviral gene transfer of Smad7 prevents injury-induced epithelial-mesenchymal transition of lens epithelium in mice. Lab. Invest. 84, 1259–1270. Salogni, L., Musso, T., Bosisio, D., Mirolo, M., Jala, V. R., Haribabu, B., Locati, M., and Sozzani, S. (2009). Activin A induces dendritic cell migration through the polarized release of CXC chemokine ligands 12 and 14. Blood 113, 5848–5856. Scutera, S., Riboldi, E., Daniele, R., Elia, A. R., Fraone, T., Castagnoli, C., Giovarelli, M., Musso, T., and Sozzani, S. (2008). Production and function of activin A in human dendritic cells. Eur. Cytokine Netw. 19, 60–68. Stupack, D. G., Cho, S. Y., and Klemke, R. L. (2000). Molecular signaling mechanisms of cell migration and invasion. Immunol. Res. 21, 83–88. Swart, G. W., Lunter, P. C., Kilsdonk, J. W., and Kempen, L. C. (2005). Activated leukocyte cell adhesion molecule (ALCAM/CD166): signaling at the divide of melanoma cell clustering and cell migration? Cancer Metastasis Rev. 24, 223–236. Talmadge, J. E. (2008). Follistatin as an inhibitor of experimental metastasis. Clin. Cancer Res. 14, 624–626. Tsuchida, K., Nakatani, M., Hitachi, K., Uezumi, A., Sunada, Y., Ageta, H., and Inokuchi, K. (2009). Activin signaling as an emerging target for therapeutic interventions. Cell Commun. Signal. 7, 15. Valcourt, U., Kowanetz, M., Niimi, H., Heldin, C. H., and Moustakas, A. (2005). TGFbeta and the Smad signaling pathway support transcriptomic reprogramming during epithelial-mesenchymal cell transition. Mol. Biol. Cell 16, 1987–2002. Wu, Y., Zhang, X., Salmon, M., Lin, X., and Zehner, Z. E. (2007). TGFbeta1 regulation of vimentin gene expression during differentiation of the C2C12 skeletal myogenic cell line requires Smads, AP-1 and Sp1 family members. Biochim. Biophys. Acta 1773, 427–439.
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Xia, Y., and Schneyer, A. L. (2009). The biology of activin: recent advances in structure, regulation and function. J. Endocrinol. 202, 1–12. Xu, J., Lamouille, S., and Derynck, R. (2009). TGF-beta-induced epithelial to mesenchymal transition. Cell Res. 19, 156–172. Yamashita, N., Nakajima, T., Takahashi, H., Kaneoka, H., Mizushima, Y., and Sakane, T. (1993). Effects of activin A on IgE synthesis and cytokine production by human peripheral mononuclear cells. Clin. Exp. Immunol. 94, 214–219. Yoneyama, H., Matsuno, K., and Matsushimaa, K. (2005). Migration of dendritic cells. Int. J. Hematol. 81, 204–207. Yu, E. W., Dolter, K. E., Shao, L. E., and Yu, J. (1998). Suppression of IL-6 biological activities by activin A and implications for inflammatory arthropathies. Clin. Exp. Immunol. 112, 126–132. Yu, J., Shao, L. E., Frigon, N. L., Jr., Lofgren, J., and Schwall, R. (1996). Induced expression of the new cytokine, activin A, in human monocytes: inhibition by glucocorticoids and retinoic acid. Immunology 88, 368–374. Zavadil, J., and Bottinger, E. P. (2005). TGF-beta and epithelial-to-mesenchymal transitions. Oncogene 24, 5764–5774. Zavadil, J., Cermak, L., Soto-Nieves, N., and Bottinger, E. P. (2004). Integration of TGFbeta/Smad and Jagged1/Notch signalling in epithelial-to-mesenchymal transition. EMBO J. 23, 1155–1165. Zhang, L., Deng, M., Parthasarathy, R., Wang, L., Mongan, M., Molkentin, J. D., Zheng, Y., and Xia, Y. (2005). MEKK1 transduces activin signals in keratinocytes to induce actin stress fiber formation and migration. Mol. Cell. Biol. 25, 60–65.
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The Synthesis and Secretion of Inhibins Kelly L. Walton, Yogeshwar Makanji, David M. Robertson, and Craig A. Harrison Contents I. Introduction II. Inhibin Expression and Regulation A. Gonadotrophins B. Additional proteins capable of regulating inhibin transcription C. Epigenetic regulation of inhibin expression D. Posttranscriptional regulation of inhibin expression III. Expression Profile of Inhibin in Human Tissues A. Cyclic inhibin expression in females B. Inhibin expression in males C. Expression of inhibins in the HPG axis D. Extragonadol expression of inhibins IV. Inhibin Assembly A. Proteolytic processing B. Posttranslational modifications C. Chaperones V. Circulating Inhibin Forms A. Circulating inhibin forms in women B. Ovarian cancer and inhibins C. Circulating inhibin B forms in men VI. Concluding Remarks References
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Abstract Inhibin A and B, dimeric glycoproteins comprising an a- and b(A/B)-subunit, negatively regulate follicle stimulating hormone (FSH) synthesis by the pituitary. The expression of a- and b-subunits within Sertoli cells of the testis and granulosa cells of the ovary is controlled by a range of transcription factors, including CREB, SP-1, Smads, and GATA factors. The inhibin a- and b-subunits are synthesized as precursor molecules consisting of an N-terminal propeptide Prince Henry’s Institute of Medical Research, Clayton, Victoria, Australia Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00008-1
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and a C-terminal mature domain. Recently, we showed that hydrophobic residues within the propeptides of the a- and b-subunits interact noncovalently with their mature domains, maintaining the molecules in a conformation competent for dimerization. Dimeric precursors are cleaved by proprotein convertases and mature inhibins are secreted from the cell noncovalently associated with their propeptides. Propeptides may increase the half-life of inhibin A and B in circulation, but they are readily displaced in the presence of the high-affinity receptors, betaglycan, and ActRII. ß 2011 Elsevier Inc.
I. Introduction Inhibin A and inhibin B are heterodimeric glycoproteins that regulate mammalian reproduction. The active heterodimers comprise an a-subunit and either a bA-(inhibin A) or bB-subunit (inhibin B). Homo- or heterodimerization of the b-subunits gives rise to the structurally related proteins, activin A (bA-subunit dimer), activin B (bB-subunit dimer), and activin AB (bA/bB dimer). Both inhibin and activin are members of the transforming growth factor-b (TGF-b) superfamily. Inhibins are produced by the Sertoli cells of the testis (Steinberger, 1979) and granulosa cells of the ovarian follicles (Woodruff et al., 1987), and act in an endocrine manner to suppress follicle-stimulating hormone (FSH) secretion from the anterior pituitary via a negative feedback loop. Inhibin regulation of FSH release occurs in a cyclic-dependent manner in females and a tonic pattern in males (Woodruff et al., 1996). Within the gonads, inhibins regulate gametogenesis and act as tumor suppressors. Inhibins exert their biological actions by antagonizing the actions of activin A and B. Activin signaling commences with binding to a type II receptor on the surface of target cells, resulting in a conformational change (Groppe et al., 2008; Hart et al., 2002). The change in conformation leads to the recruitment and phosphorylation of a type I receptor (Wrana et al., 1994) and subsequent initiation of an intracellular signaling cascade involving Smad transcription factors (Derynck et al., 1998). Inhibins, in conjunction with the coreceptor betaglycan, form a high-affinity complex with the activin type II receptor (Lewis et al., 2000). This interaction inhibits activin access to its type II receptors and blocks the intracellular signaling cascade. In this chapter, we will initially describe the transcriptional and posttranscriptional processes that regulate the expression of the inhibin a- and b-subunits. Subsequently, we will summarize our recent studies, which have characterized the biosynthetic pathway that governs the assembly and secretion of inhibin A and B.
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II. Inhibin Expression and Regulation Inhibin synthesis commences with transcription of the genes coding for the a-, bA-, and bB-subunits. In humans, the inhibin a-subunit gene is located on chromosome 2 (2q33-q36; Barton et al., 1989) and is highly conserved among all species (Mason et al., 1986). The human inhibin a-subunit gene is comprised of two exons separated by a 1.7 kbp intron (Stewart et al., 1986). The bA- and bB-subunit genes are located on chromosome 7 (7p15-p13) and 2 (2cen-q13), respectively (Mason et al., 1986). The human bA-gene comprises three exons and a 2.6 kbp intron region. The human bB-subunit consists of two exons separated by a 2.5 kbp intron (Mason et al., 1986). Two bB-subunit mRNAs of 3.8 and 4.8 kbp have been observed in human tissues (Dykema and Mayo, 1994; Feng et al., 1995). These transcripts arise from the different promoter elements within the 50 flanking region of the gene. Inhibin subunit expression is tightly controlled by promoter elements in the 50 untranslated regions (UTRs) of the genes (Fig. 8.1). Sequence variations in the 50 UTRs regulate the differential expression of the inhibin genes. Expression of the inhibin a- and bA-subunits is initiated by RNA polymerases at conventional TATA boxes (Feng et al., 1989; Tanimoto et al., 1991). In contrast, the bB-subunit lacks the TATA element (and CAAT-like sequences), suggesting that bB-mRNA transcription is initiated via a different mechanism. Transcription of the bB-subunit likely involves the GC-rich regions of the promoter, and specificity protein 1 (SP-1) and activator protein-2 (AP-2) transcription factors (Dykema and Mayo, 1994; Mason et al., 1989). The SP-1 transcription factor also appears to modulate the expression of the a- and bA-subunits. Binding sites for GATA factors have also been identified in the inhibin a- and bA-subunits (Fig. 8.1), and supporting experiments have demonstrated that these factors are transcriptional activators of inhibins (Robert et al., 2006; Tremblay and Viger, 2001).
A. Gonadotrophins Transcriptional activation of inhibins in the ovary and testis is modulated by pituitary gonadotrophins, FSH and luteinizing hormone (LH; Bicsak et al., 1986; Suzuki et al., 1987; Tsonis et al., 1987; Woodruff et al., 1987; Ying et al., 1987; Zhang et al., 1987). The gonadotrophins promote inhibin expression by activation of G protein-coupled receptors, resulting in an increase in intracellular cAMP levels via activation of adenylyl cylase. The increase in cAMP activates the protein kinase A (PKA) pathway, leading to the phosphorylation of cAMP responsive element binding protein (CREB;
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A Chromosome location:2q33-q36 220,436,994
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GATA AP-1 AP-1 AP-2 GATA SF-1 SMAD 3/4 CREB TRE TRE AP-2 GATA SF-1 SBE CRE GATA B Chromosome location:7p15-p13 41,742,06 INHBA 5⬘U
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Figure 8.1 Model of the inhibin a- (A), bA- (B), and bB-subunit (C) genes. The mRNA transcripts for each of the subunits are mapped (second line), and important regulatory elements within the 50 UTRs are noted, including GATA binding sites, smad-binding elements (SBE), CAMP-responsive elements (CRE), and 12-O-tetradecanoyl phorbol 13-acetate responsive elements (TREs). Transcriptional activators that act as these sites are also marked; cAMP binding protein (CREB), steriodogenic factor-1 (SF-1), liver receptor homolog 1 (LRH-1), and activator proteins-1 and 2 (AP-1 and -2).
Mukherjee et al., 1996; Pei et al., 1991). Phosphorylation of CREB stimulates inhibin transcription via a CREB-mediated interaction with the CRE in the promoter (Ardekani et al., 1998; Pei et al., 1991; Tanimoto et al., 1996).
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Transcription factors, inducible cAMP early repressor (ICER) protein (Mukherjee et al., 1998), and CCAAT/enhancer binding protein b (C/EBPb; Burkart et al., 2005) attenuate the stimulatory effects of cAMP by competitively binding to the CRE in the promoter. Suppression of FSH secretion attenuates the transcription of the inhibin subunits. Gonadotrophin-releasing hormone (GnRH) from the hypothalamus, testosterone from the testis, estradiol and progesterone from the ovaries, androgens and corticosteroids (Suzuki et al., 1987; Tsonis et al., 1987), and gonadal inhibins, local activins, and follistatin, all regulate the synthesis of gonadotrophins from the anterior pituitary. GnRH secretions from the hypothalamus modulate the expression of FSH and LH by the gonadotropes (Sealfon et al., 1997; Shacham et al., 2001). Inhibins produced by the gonads act in an endocrine manner to control FSH production at the pituitary. Gonadal inhibins downregulate FSH production at the pituitary by antagonizing the actions of locally produced activins (Attardi and Miklos, 1990; Attardi et al., 1992; Carroll et al., 1989; Corrigan et al., 1991). The activin antagonist, follistatin (FST), also negatively regulates FSH secretion (Kaiser et al., 1992). Evidence supports that activin A also stimulates LH production in gonadotrophic cells, at both the mRNA and protein level (Coss et al., 2005; Yamada et al., 2004). Importantly, inhibin suppression of FSH release at the pituitary provides a negative feedback loop to tightly control inhibin secretions during folliculogenesis in the ovary and spermatogenesis in the testis.
B. Additional proteins capable of regulating inhibin transcription Inhibin A and B appear to be differentially regulated within the ovary. Unlike the a- and bA-subunits, transcription of the inhibin bB-subunit is not regulated by cAMP responsive elements (Dykema and Mayo, 1994). Expression of the bB-subunit is, however, regulated by various members of the TGF-b superfamily, including activin A (Eramaa et al., 1995), BMP-2 ( Jaatinen et al., 2002), TGF-b1 and -b2 (Eramaa and Ritvos, 1996), GDF-9 (Kaivo-Oja et al., 2003), and BMP-15 (Edwards et al., 2008). The stimulatory effects of the TGF-b ligands can be mimicked by overexpressing the intracellular signaling components of these pathways, including Smad 1 and 2 (Bondestam et al., 2002; Kaivo-Oja et al., 2003). The findings suggest that the TGF-b ligands trigger Smad-mediated transcriptional regulation of the inhibin bB-subunit. It is unlikely however, that the responsive Smads are interacting directly with the bB-subunit promoter to regulate transcription, as the bB-promoter lacks a consensus Smad-binding element (SBE; ACAGACA; Dennler et al., 1998; Zawel et al., 1998). In contrast, the inhibin a-promoter does contain a SBE (Fig. 8.1), suggesting that transcription of this subunit may be directly mediated by Smads. The SBE appears
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to be specifically responsive to TGF-b/activin signaling pathways (Dennler et al., 1998). In support, BMP-6 is capable of upregulating mRNA expression of both the inhibin bA- and bB-subunits, but has no effect on the inhibin a-subunit (Shi et al., 2009). BMP-6 also increases the FSH receptor mRNA levels (by approximately threefold), suggesting that the increase in subunit expression could be attributed to an increased responsiveness to FSH in these cells. In summary, the regulation of inhibin synthesis at the transcriptional level is complex, requiring the stimulation of various transcription factors, which target specific regions of the promoters of the individual subunits. Thus, the differential production of inhibin and activins can in part, be attributed to the tissue-specific expression of the transcription factors and sequence variations within the promoter regions of the subunit genes that bind these activators.
C. Epigenetic regulation of inhibin expression Epigenetic regulation, or the modification of gene activation, is another means of transcriptional control of the inhibin subunit genes. In most cases, epigenetic modifications alter the interaction between transcriptional activators and target promoters, impacting gene synthesis. One such modification is DNA methylation. The human inhibin a-subunit promoter contains seven potential CpG methylation sites (149 to 284, relative to ATG at position 1). Four of these sites are unique to humans (CpG1, CpG2, CpG5, and CpG6). A study has shown that the inhibin a-subunit promoter is hypermethylated in both low- and high-grade prostate cancer samples relative to controls (Schmitt et al., 2002). Hypermethylation is predicted to cause transcriptional repression, resulting in loss of the inhibin a-subunit mRNA in these tissues. Moreover, extensive methylation of the inhibin a-subunit promoter in prostate cancer cells lines (LNCap, DU145, and PC3) also correlates with a lack of inhibin a-mRNA (Balanathan et al., 2004). Supporting studies have shown that the inhibin a-subunit acts as a tumor suppressor in gonadal and adrenal tissues (Matzuk et al., 1992, 1994). Histone modification has also been associated with the transcriptional regulation of the inhibin subunit genes. Salvador et al. demonstrated that FSH stimulates both PKA-mediated phosphorylation and acetylation of histone H3 in primary rat granulosa cells (Salvador et al., 2001). The modifications to histone H3 appear to increase its association with the inhibin a-promoter, resulting in increased transcription. Transcriptional activators, PKA and SF-1, also increase histone H4 acetylation associated with the inhibin a-promoter (Ito et al., 2000). mRNA expression of the bA-subunit is also regulated by histone acetylation and DNA promoter methylation in lung adenocarcimona cell lines (Seder et al., 2009). Cotreatment of cells with 5-aza-20 -deoxycytidine (demethylation agent) and
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trichostatin A (histone deacetylase inhibitor) substantially enhances expression of the bA-subunit (up to 86-fold). These results suggest that DNA and histone modifications negatively regulate the transcription of the inhibin bA-subunit.
D. Posttranscriptional regulation of inhibin expression MicroRNAs (miRNAs) are an extensive class of evolutionarily conserved 18–24 nucleotide noncoding RNAs that posttranscriptionally regulate gene expression. This is achieved by imperfectly base-pairing with specific sequences in the 30 UTRs of target mRNAs, thus inhibiting their translation (Ambros, 2001); hence, miRNAs typically act as negative regulators of protein expression. Little is known about the role of miRNAs in the regulation of TGF-b ligand expression. Using bioinformatic analysis, we identified miRNAs predicted to target the inhibin subunit genes. The 30 UTR of the inhibin bB-subunit (1.9 kb) is significantly longer than the corresponding regions of the inhibin bA- (0.64 kb) and a-subunits (0.18 kb) and is predicted to have conserved binding sites for numerous miRNAs, including mir-106, mir129, and mir-148. In contrast, only two conserved target sites for miR-205 and miR-135 were identified in the 30 UTR of the bA-subunit. There are no predicted conserved miRNA binding sites in the 30 UTR of the human inhibin a-subunit. Interestingly, miR-129 is one of the most highly expressed miRNAs in the seminiferous tubule of the testis (unpublished observation), where it may play a role in the regulation of inhibin bBsubunit translation. Currently, we are exploring whether the other identified miRNAs are expressed in Sertoli and granulosa cells.
III. Expression Profile of Inhibin in Human Tissues Transcriptional regulation controls the tissue-specific expression of inhibin mRNAs and proteins. The gonads are the main production sites of inhibins, but extragonadal expression has been observed in several tissues (Table 8.1).
A. Cyclic inhibin expression in females Expression of the inhibin subunit mRNAs occurs in a cyclic-dependent manner in females. All of the subunit mRNAs are expressed in the ovary, but there is 10-fold more a- than bA- or bB-subunit mRNA (Mason et al., 1985; Meunier et al., 1988; Woodruff et al., 1987). The smaller antral
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Table 8.1 Tissue distribution of activin and inhibin subunits, betaglycan (BG), activin receptors, and follistatin (FST) mRNA in adult tissues
Tissue
Ovary Placenta Uterus/decidua Oocytes Small antral Granulosa follicles cells Theca cells Granulosa Large cells dominant follicles Theca cells Testis Sertoli cells Leydig cells Prostate Brain Anterior pituitary Gonadotropes Adrenal Bone marrow Breast Spleen Heart Lung Thymus Skeletal muscle Kidney Pancreas Liver/hepatocytes
Act Act bA bB a BG RIIA RIIB ALK4 ALK7 FST
þ þ þ þ þ
þ þ ? þ þ
þ þ þ þ þ
þ þ þ þ þ
þ þ þ þ þ
þ ? ? þ ?
? þ
þ þ þ ?
þ þ þ þ þ þ þ
þ
þ
? þ
þ þ
? ?
þ þ þ þ þ ? þ þ þ þ þ þ þ þ
þ þ ? þ þ þ þ ? ? þ ? þ þ þ þ ? þ þ
þ þ ? þ þ þ ? ? ? þ ? þ ? þ ? ? þ
þ þ ? þ þ ? þ ? ? þ þ þ þ þ þ þ ? þ
? ? ? þ ? þ þ þ
þ þ ? þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ
þ þ þ þ þ þ þ þ þ þ þ þ þ þ
þ þ þ þ
þ þ þ þ þ þ þ þ þ þ ? ? þ ? ? ? ? þ
þ þ ? þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ
Adapted from Alexander et al. (1996), Anderson et al. (2002), Bondestam et al. (2002), Casagrandi et al. (2003), Chen and Johnson (1996), DePaolo et al. (1991), Di Loreto et al. (1999), Haddad et al. (1994), He et al. (1993), Jeruss et al. (2003), Jones et al. (2002b), Knight and Glister (2006), La Rosa et al. (1999, 2004), MacConell et al. (2002), Martins da Silva et al. (2004), Mathews and Vale (1991), Matsuzaki et al. (1993), Ohga et al. (2000), Pangas et al. (2002), Penabad et al. (1996), Reis et al. (2004), Roberts et al. (1993, 1994), Sidis et al. (1998), Tuuri et al. (1994), van Schaik et al. (2000)Wada et al. (1996), Walsh et al. (2007), Wang and Tsang (2007), Wang et al. (2006), Yndestad et al. (2004), Yu et al. (1994) and expression profiles from http://www.ncbi.nlm.nih.gov/sites/entrez?db¼unigene of Hs.583348INHBA, Hs.1735-INHBB, Hs.470174-ACVR2A, Hs.174273-ACVR2B, Hs.4389818-ACVR1B, Hs.562901-ACVR1C, and Hs.9914-FST.
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follicles express mainly the bB-subunit, whereas the dominant follicles and corpus luteum express mainly the bA-subunit (Roberts et al., 1993; Welt et al., 2001). Inhibin a-subunit expression is consistent in follicles of all sizes. Although b-subunit expression has been identified predominantly in the granulosa cells of the developing follicle, expression has also been observed in the thecal cells of human dominant follicles (Roberts et al., 1993). The varying b-subunit expression accounts for the different inhibin isoforms secreted by the ovary during the reproductive cycle. The small antral follicles produce mainly inhibin B, while the dominant follicles and corpus luteum secrete inhibin A (Welt and Schneyer, 2001; Welt et al., 2001). Betaglycan and activin type II receptor mRNAs have been detected within the theca and granulosa cells and oocytes during all stages of development (Drummond et al., 2002). The activin antagonist, follistatin, is expressed by the granulosa cells of all sizes during folliculogenesis. The controlled secretions of FSH and LH regulate the production of the various inhibin isoforms in the ovary (Groome et al., 1996). Inhibin B rises across the luteal–follicular transition, reaching a peak in the mid-follicular phase and a second peak on the day after the LH surge. Declining inhibin B levels in the luteal–follicular transition correlate with an increase in FSH (Burger et al., 1998b; Klein et al., 1996; Reame et al., 1998; Welt et al., 2003). Inhibin A begins to rise in the late follicular phase, peaking at ovulation and is at its maximal level in the mid-luteal phase (Groome et al., 1996; Welt, 2004).
B. Inhibin expression in males In males, FSH drives the tonic production of inhibin B in the Sertoli cells of the testis (Grootenhuis et al., 1990a,b; Hancock et al., 1992; Illingworth et al., 1996; Marchetti et al., 2003). The bB-subunit mRNA has been detected in the Sertoli, Leydig, and germ cells of adult human testes tissue (Buzzard et al., 2004; Majdic et al., 1997; Marchetti et al., 2003). The inhibin a-subunit mRNA is produced predominantly by the Sertoli cells (Bicsak et al., 1987) of the testis, but has also been identified within the Leydig cells (Risbridger et al., 1989). Inhibin a-subunit mRNA levels are maximal between stages XIII–I and minimal between stages VII–VIII of the rat seminiferous cycle (Bhasin et al., 1989; Kaipia et al., 1991). Consequently, inhibin B secretion is highest from stage IX–I and lowest at stage VII (Okuma et al., 2006). Although the quantification of activin B expression in the testis has been limited by the available detection methods, high levels of activin B have been identified in seminal plasma fluid of adult human males (Ludlow et al., 2009), and activin B is a proliferative factor for Sertoli cells in vitro (Mather et al., 1990). Adult males lack any circulating inhibin A, but do synthesize activin A, as supported by studies showing that activin A can induce Sertoli cell proliferation (Boitani et al., 1995).
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The bA-subunit mRNA has been localized to the Sertoli and Leydig cells of the adult human testis (Vliegen et al., 1993) and is expressed in stages VII–XII.
C. Expression of inhibins in the HPG axis Inhibins are key regulators of the hypothalamic pituitary gonadal axis (HPG), and both b-subunits are detectable in the pituitary gonadotropes (Roberts et al., 1992). The bA-subunit is also expressed in somatropes and lactotropes, whereas the bB-subunit is expressed in thyrotropes (Uccella et al., 2000). The inhibin a-subunit mRNA is also expressed in the rat pituitary (Meunier et al., 1988), although its precise location has not been described. It is proposed that activin B may act in an autocrine manner at the pituitary to regulate FSH secretion, as it has been shown that neutralization of activin B activity inhibits FSH secretion in cultured rat pituitary cells (Corrigan et al., 1991).
D. Extragonadol expression of inhibins Extragonadal expression of the inhibin subunit mRNAs have been described in a variety of adult human tissues, including the placenta, endometrium, pituitary gland, prostate (Thomas et al., 1998), breast (Di Loreto et al., 1999), adrenal gland (Spencer et al., 1990; Spencer et al., 1992), lung, liver, and bone (Centrella et al., 1994; see Table 8.1). In addition, the target activin receptors and betaglycan are also expressed in these tissues, suggesting that inhibin could act in an autocrine or paracrine manner to regulate activin signaling at these sites. Thus, it is likely that activin and inhibin action is determined by the differential tissue expression of target receptors and accessory binding proteins (Table 8.1). For example, the endometrium produces large quantities of the inhibin bA- and bB-subunit mRNAs, resulting in high expression of activins during the secretory phase ( Jones et al., 2000; Leung et al., 1998). In contrast, the inhibin a-subunit is only moderately expressed in the glandular and surface epithelium, such that inhibin A and B protein levels are 1000-fold lower than that of activins ( Jones et al., 2006; Petraglia et al., 1998). At the end of the menstrual cycle, during decidualization and in early pregnancy, the inhibin a-subunit mRNA expression shifts from epithelial to stromal cells ( Jones et al., 2000). Decidualized human endometrial cells in culture respond to activin A treatment by elevating matrix metalloproteinase secretion, and inhibin A blocks this activin-mediated response ( Jones et al., 2006). Expression of the inhibin a-subunit, bA- and bB-subunit mRNAs increases throughout pregnancy, reaching maximal levels in the third trimester (Petraglia et al., 1990). Inhibin A plays some important roles in placentation and in pregnancy. In contrast to low levels of inhibin A produced by the nonpregnant uterus, the syncytiotrophoblasts cells of
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the placenta produce abundant inhibin A (Qu and Thomas, 1995), together with betaglycan (Ciarmela et al., 2003; Jones et al., 2002a). In the placenta, inhibin is a potent antagonist of activin-mediated steroidogenesis and hCG production by syncytiotrophoblasts ( Jones et al., 2006; Petraglia et al., 1989). Throughout the gestation period, levels of activin A and inhibin A continue to rise up until parturition (Muttukrishna et al., 1995). The precise role of inhibins in the placenta is not understood; however, in several placental diseases, inhibin A is implicated. The controlled expression of inhibins and activin in female reproduction is essential to the homeostasis of this system. Aberrant inhibin/activin expression has been linked to deleterious pregnancy outcomes, ranging from alterations in placental mass (Casagrandi et al., 2003; Muttukrishna et al., 2004), to preeclampsia (Bersinger et al., 2003) and miscarriage (Wallace et al., 2004). Elevated levels of serum inhibin A in the second trimester of pregnancy is indicative of fetal Down’s syndrome, and is utilized today as a marker for this disease in combination with other factors, including hCG and a-fetoprotein (Malone et al., 2005; Wallace et al., 1996). Inclusion of the measurement of maternal serum inhibin A and activin A along with other tests in the second trimester of pregnancy may improve the predictive efficacy of early onset preeclampsia screening (Ay et al., 2005; Florio et al., 2004; Madazli et al., 2005; Spencer et al., 2006). Many of the studies examining the tissue and cellular localization of inhibins are supported by experiments associating the aberrant expression of inhibins/activins with cancerous states. For example, the significance of inhibin production by the adrenals (Spencer et al., 1990, 1992) became apparent when Matzuk and colleagues (Matzuk et al., 1994) demonstrated that inhibin a-subunit knockout mice developed adrenocortical tumors with 99% penetrance after gonadectomy. Moreover, the inhibin subunits are expressed in both the normal human prostate and prostate cancer cells, but loss of the inhibin a-subunit is consistent with prostate cancer malignancies (Mellor et al., 1998; Thomas et al., 1998). Similarly, the mammary gland of the breast expresses the inhibin a-, bA-, and bB-subunits and secretes activin A, and inhibin A and B (Di Loreto et al., 1999), but in breast cancer, the levels of bA-mRNA and activin A protein are notably higher (Reis et al., 2002). Thus, characterization of the expression profiles of the inhibin transcripts and proteins is a valuable tool for monitoring the homeostasis of human tissues.
IV. Inhibin Assembly Following transcription, the inhibin a-subunit is translated as a large precursor protein with a 232-amino acid pro-region and a 134-amino acid mature region (termed aC; Mason et al., 1996; Stewart et al., 1986).
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A secondary dibasic arginine processing site within the pro-region of the a-subunit further divides this domain into pro- (43 amino acids) and aNregions (171 amino acids). Characteristic of TGF-b family members, the inhibin a-subunit contains seven conserved cysteine residues in the mature domain, forming a cysteine knot motif crucial for monomer and dimer assembly. The bA-subunit precursor comprises a 290-amino acid proregion and a 116-amino acid mature bA region (Mason et al., 1996). Similarly, the bB-subunit comprises a 264-amino acid pro-region and a 115-amino acid mature region. Within the mature domains of the bA- and bB-subunits, nine cysteine residues are required for inter- and intramolecular disulfide bond formation. The N-terminal signal peptide of the monomeric inhibin subunits targets them to the endoplasmic reticulum (ER; Kingsley, 1994; Massague et al., 1990). Within the ER, the prodomains of the monomeric subunits are necessary for the correct folding, disulfide bond formation, export, and biological activity of inhibin A and B (Gray and Mason, 1990; Fig. 8.2). Recently, we performed structure/function analyses of the prodomains of the inhibin a- and bA-subunits to identify the key residues that promote covalent dimerization and drive inhibin assembly (Gray and Mason, 1990; Walton et al., 2009). Within the a-subunit, we utilized site-directed mutagenesis to identify three hydrophobic residues near the N-terminus of the prodomain (Leu30, Phe37, Leu41) that interact noncovalently with hydrophobic residues in the mature domain (Phe271, Ile280, Pro283, Leu338, and Val340) to maintain the molecule in a conformation competent for dimerization (Walton et al., 2009). Protein structure prediction (QuickPhyre, Imperial College, London) indicated that the N-terminal region of the a-subunit prodomain encompasses an a-helix, and modeling (PyMOL Molecular Graphics System) predicted that the helical domain was amphipathic. Hydrophobic a-subunit prodomain residues (Leu30, Phe37, Leu41), which bind to the mature a-subunit, formed a contiguous epitope on one surface of the a-helix. Interestingly, sequence alignment indicated that the a-helical region is conserved in the prodomains of other TGF-b superfamily members (Fig. 8.3). Correspondingly, hydrophobic residues within the bA-subunit prodomain, including Ile62 and Leu66, were shown to interact noncovalently with Phe329, Ile338, Pro341, Met398, and Met400 in the mature domain to ensure correct folding of the molecule. The inhibin a- and bAsubunits then interact noncovalently at two sites: the first close to the cysteine knot motif in the mature domains and the second near the C-terminus of the prodomains (Walton et al., 2010). Noncovalent dimerization of the mature domains facilitates interchain disulfide bond formation between Cys327 in the inhibin a-subunit and Cys390 in the bA-subunit (Husken-Hindi et al., 1994). Dimerization of the a- and bA-prodomains is predicted to stabilize the complex (Walton et al., 2010). Significantly,
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Inhibin Synthesis and Secretion
A SP
Prodomain
Mature
B
a-Subunit
b-Subunit
C 1
Prodomain
2
Mature inhibin
Granulosa/Sertoli cell D
E
ActRII/IIB
Betaglycan
Pituitary cell
Figure 8.2 Model for inhibin/activin synthesis and secretion. (A) Inhibins are synthesized from large precursor proteins, comprising an N-terminal prodomain and a C-terminal mature domain. (B, C) Hydrophobic residues within the N-terminal portion of the prodomain (region 1) and regions of the mature domain interact noncovalently, maintaining the molecule in a conformation competent for dimerization. The prodomains of the inhibin subunits also dimerize via a C-terminal noncovalent interaction (region 2). The prodomains are cleaved from the mature dimer by proprotein convertases, but remain noncovalently associated with the mature growth factor. (D) The inhibin complex is secreted from the cell. (E) High-affinity interactions with binding partners on the surface of target cells cause the prodomain to dissociate from the mature ligand. Adapted from Walton et al. (2009, 2010).
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Figure 8.3 Sequence alignment of the N-terminus of the inhibin prodomains. A region at the N-terminus of the TGF-b prodomains forms an amphipathic alpha helix. The helical nature of this region is conserved in most TGF-b ligands (gray). Hydrophobic residues (bold) in this region interact with residues in the mature ligand to mediate dimer formation. Cationic residues in this region (underlined) of the TGF-b1 prodomain have been implicated in binding to LTBP-1.
disruption of the noncovalent interactions between the pro- and mature domains of the inhibin a- and bA-subunits completely abrogates inhibin A expression (Walton et al., 2009).
A. Proteolytic processing Following dimerization, the inhibin a- and b-subunit prodomains are enzymatically cleaved from the mature dimers at consensus RXXR sites (Kingsley, 1994; Massague et al., 1990; Molloy et al., 1999). The inhibin a-subunit possesses two proteolytic target sites, the first (RLPR, separating the pro- and aN-regions), and the second (RARR) between the aN and aC (mature region). Both of these proteolytic sites are predicted target sites for the proprotein convertase, furin. The bA- and bB-subunits possess only single cleavage sites between their pro- and mature domains.
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The bA-subunit contains the consensus furin motif, RRRR, yet the bBsubunit cleavage site is instead RIRKR. Although the bB-subunit cleavage site does not contain the exact furin cleavage motif, it still satisfies the sequence requirements for furin cleavage (Nakayama, 1997). The requirement for furin processing is emphasized in experiments that show that activin A can drive furin mRNA expression (Antenos et al., 2008). Antenos et al. demonstrated that release of the inhibin and activin B dimers from their respective prodomains is mediated by furin.
B. Posttranslational modifications Two N-linked glycosylation sites (Asn146 and Asn302; human numbering) are present in the inhibin a-subunit of most species. However, the human a-subunit has three N-linked glycosylation sites; at Asn146, Asn268, and Asn302, and this difference is responsible for the molecular mass heterogeneity observed with human inhibin A and B forms. Asn268 is always glycosylated (giving rise to 31 kDa inhibin A or B), whereas Asn302 is differentially glycosylated (34 kDa inhibin A or B; Makanji et al., 2007; Mason et al., 1996). The functional consequence of this additional glycosylation site will be discussed elsewhere in this issue. The bA-subunit has only one glycosylation site in the prodomain of the protein (Asn165). A single glycosylation site is also predicted in the prodomain of the bB-subunit at Asn93 (NetNGlyc 1.0); however, this modification has not been experimentally validated. N-linked glycosylation of the inhibin a-subunit is required for inhibin A assembly and secretion (Antenos et al., 2007). A single glycan moiety at Asn268 ensures proper folding of the polypeptide chains and directs the overall production of inhibin dimers. Perturbation of the glycans results in intracellular accumulation of the protein, and alterations in the ratio of inhibin/activin secreted. Interestingly, it was found that inhibin A secretion could be enhanced by approximately fourfold with the addition of a glycosylation site in the bA-subunit (Phe327, analogous to Asn268 in the inhibin a-subunit). The bioactivity of inhibin is dependent on its glycosylation state (Makanji et al., 2007); the monoglycosylated form of inhibin (Asn268) is more bioactive than the diglycosylated variant (Asn268 and Asn302). The reduced bioactivity can be partly attributed to diglycosylated inhibin A having lower affinity for its coreceptor betaglycan. These findings support earlier studies that emphasize the requirement for glycosylation in TGF-b secretion and bioactivity (Brunner et al., 1992). Some members within the TGF-b superfamily, including GDF-9 and BMP-15, are phosphorylated within their mature domains (McMahon et al., 2008; Saito et al., 2008; Tibaldi et al., 2010). Importantly, phosphorylation is required for the biological activity of the GDF-9 and BMP-15 mature growth factors (Matzuk et al., 1994; McMahon et al., 2008; Saito et al., 2008; Tibaldi et al., 2010). Phosphorylation sites are predicted within the inhibin subunits
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(NetPhos version 2.0); however, evidence suggests that the bA-subunit is not phosphorylated (McMahon et al., 2008). Additional studies are required to characterize the phosphorylation status of the inhibin A and B forms.
C. Chaperones As previously discussed, the prodomains of the inhibin a- and b-subunits mediate the assembly of inhibin and activin dimers (Walton et al., 2009). The prodomains of the TGF-b isoforms (TGF-b1, -b2, -b3) play a similar role, but they also interact with intracellular chaperone molecules, termed latent TGF-b binding proteins (LTBPs; Brunner et al., 1989; Miyazono et al., 1988; Wakefield et al., 1988). Recently, we identified the ionic residues (Arg45, Arg50, Lys56, and Arg58) on the TGF-b1 prodomain, which form the binding site for LTBPs (Walton et al., 2010). Following binding, disulfide exchange occurs between a pair of cysteines in LTBPs and Cys33 in the prodomain of TGF-b1 (Gleizes et al., 1996; Rifkin, 2005; Saharinen et al., 1996). The resulting large latent complex, comprising the mature growth factor, prodomains, and bound LTBP, is secreted from the cell. LTBP targets TGF-b to fibrillin microfibrils within the extracellular matrix. This interaction regulates signaling by shielding the receptor interacting epitopes on the mature ligand (Bottinger et al., 1996; De Crescenzo et al., 2001). An activation mechanism, mediated by integrins, thrombospodin-1, or proteases, is required to release the mature growth factor for signaling (Annes et al., 2003; Yang et al., 2007; Young and Murphy-Ullrich, 2004). Given that the LTBP-1 binding motif is poorly conserved in the inhibin a- and b-subunits, it is unlikely that inhibin A and B interact with these chaperones. Intriguingly, however, the cysteine residue (Cys33 in TGF-b1) that promotes covalent association between the TGF-b1 prodomain and LTBP is conserved in the inhibin a-subunit (Cys19). This cysteine can form an intramolecular disulfide bond with the mature domain, resulting in the proa/aC inhibin form. It is possible that this free cysteine may serve to promote inhibin association with an intracellular accessory binding protein during ligand assembly. We have demonstrated that mutation of this cysteine residue (C19S) significantly reduces inhibin production, and this is also marked by a loss of a 50 kDa precursor inhibin form (Walton et al., 2009). If there is an intracellular chaperone interacting with Cys19 in the prodomain of the inhibin a-subunit, however, it is not essential for inhibin folding and secretion.
V. Circulating Inhibin Forms The numerous secreted forms of inhibin (ranging from 29–100 kDa) can be attributed to the regulated expression of the processing enzymes and variations in posttranslational modifications in target tissues (Fig. 8.4). The generation of specific monoclonal antibodies for the inhibin a- and
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Inhibin Synthesis and Secretion
A
Monomeric inhibin a- and b-subunits ~ Pro
aN
aC
kDa ~52–58
~20K
~27–30
~20K
~18–21
~ ~9K ~23K
ProbA/B ~46K
~13K ~59
~13K
B
Dimeric forms of inhibin/activin A and B ~9K
~23K ProbA/B ~46K
~20K
a ~111
b
bA,B
~20K ~46K
~9K
~23K
~13K
bA,B
~20K
a ~65 bA,B
~13K
~20K Inhibin ~13K ProbA/B ~46K
~13K
ProbA/B ~46K
~13K ~13K
Activin A, B, or AB
a ~79
~13K
a ~31–34 bA,B
~26 kDa
Figure 8.4 Precursor and mature forms of inhibin/activin A and B. The molecular masses of the subunits are indicated in kDa. The cleavage (^) and potential (*) glycosylation sites are noted.
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b-subunits, and subsequent development of specific ELISAs, has facilitated the identification of the various circulating forms of inhibin (Groome, 1991; Groome and Lawrence, 1991; Groome et al., 1990, 1996; Ludlow et al., 2008, 2009; Robertson et al., 2001).
A. Circulating inhibin forms in women Several groups have characterized the molecular forms of inhibin in follicular fluid (FF) and serum/plasma of numerous species (ovine, porcine, bovine, equine, rat, and human; Table 8.2). The 31 and 34 kDa heterodimers of the aC- and b-subunits are regarded as the mature inhibin forms, whereas uncleaved and partially cleaved dimers of the a- and b-subunits are classed as high molecular weight inhibins (Fig. 8.4). Robertson and colleagues characterized the various biological forms of inhibin in human follicular fluid (hFF) and serum of women undergoing gonadotrophin stimulation as part of IVF programs (Robertson et al., 1996, 1997). The hFF samples were fractionated using a combination of inhibin aC-subunit immunoaffinity purification and reverse-phase HPLC. The various forms of inhibin were determined using ELISAs. Human follicular fluid contains inhibin aC-precursor forms (66 and 55 kDa) and mature inhibin glycoforms (31 and 34 kDa; Fig. 8.4 and Table 8.2). High molecular weight forms of inhibin (100 kDa) have also been observed in hFF (Robertson et al., 1996). Similar forms of inhibin A and B are observed in IVF serum, with the exception of a 55 kDa aC-precursor form. The 55 kDa aC-precursor is only observed for inhibin A. The 31 kDa mature form of inhibin A is also not observed in IVF serum. The findings support that inhibin A and B are differentially processed in circulation. During pregnancy, a range of molecular weight forms (31–100 kDa) of inhibin A are observed in the placenta, amniotic serum, and maternal serum (Thirunavukarasu et al., 2001, 2003). The relative abundance of the mature inhibin A forms increases from the first to third trimesters, whereas the other forms of inhibin A appear to dominate during the earlier phases of pregnancy (first and second trimesters). It is suggested that the increase in mature processed inhibin A results from the increasing presence of circulating target proteases. Detection of inhibin B during pregnancy has been limited by the sensitivity of the available assays. However, inhibin B has been detected in the amniotic fluid, where the mature inhibin B form appears to be the most abundant.
B. Ovarian cancer and inhibins Considerable attention has been given to the contribution of inhibins in the pathogenesis of ovarian cancer (Burger et al., 1996; Robertson et al., 1996). Ovarian cancers are classed as either epithelial, stromal sex cord, or germ cell.
Table 8.2
Various molecular mass (kDa) forms of inhibin A in mammalian species Mature forms
Species
Human
Biological fluid
Unknown
ProaN-aC /Prob-b
Unknown
ProaNaC/b
aN-aC /Prob-b
aNaC/b
aNaC
aCa /b
aCb /b
ProaC
aN
aC
References
– – –
– 122 –
97 97 97
– – –
66 66 66
56 56 56
– – –
36 36 36
33 – 33
29 29 –
– – –
– – 21
Robertson et al. (1996, 1997)
– –
– –
– 105
– 75
66 –
56 58
– –
36 –
– 30
29 27
– 23
21 –
Bovine
hFF IVF Serum Postmenopausal women Male plasma bFF
>120
120 (116)
108
88
65
56–58
44
–
32
25
–
–
>160
122
–
77 (?)
68
58
48 49
–
33
29
–
–
>160
110
–
53–58
49
–
34
29
–
–
Good et al. (1995)
–
–
–
88 77 –
–
Ovine
bFF (utero–ovarian venous or peripheral plasma) bFF (from estrogen active, atretic, and highly atretic follicles) bFF immunopurified and electroeluted oFF
Robertson et al. (1989), Sugino et al. (1992) Knight et al. (1989), Miyamoto et al. (1986) Ireland et al. (1994)
65
–
–
–
30
29
–
–
Sertoli cell conditioned media
–
–
–
–
–
–
45
–
32
29
–
–
Leversha et al. (1987) Grootenhuis et al. (1990a)
Rat
(continued)
Table 8.2
(continued) Mature forms
Species
Biological fluid
Porcine
Testicular homogenates Sertoli cell culture medium pFF
Equine Primate (Macaca mulatta) a b
eFF Serum (males)
Diglycosylated aC. Monoglycosylated aC.
Unknown
ProaN-aC /Prob-b
Unknown
ProaNaC/b
aN-aC /Prob-b
aNaC/b
aNaC
aCa /b
aCb /b
ProaC
aN
aC
References
–
–
–
–
–
–
–
32
–
–
–
–
–
–
–
–
44 43 –
–
29
27
23
–
Noguchi et al. (1997) Hancock et al. (1992)
–
–
–
100
80
–
55
–
–
32
–
–
–
– –
– –
90 90–100
80 –
– 56–60
56 –
40 –
– –
32 36–31
– 26–29
– –
– –
Ling et al. (1985), Miyamoto et al. (1985), Rivier et al. (1985) Moore et al. (1994) Majumdar et al. (1997), Winters and Plant (1999)
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The epithelial cancers make up 90% of the all ovarian cancers and are further characterized as serous (70%), muscinous (10–15%), endometroid (10–15%), and other subtypes. Granulosa cell tumors are present in only 5% of cases (Robertson et al., 2007). During menopause, the depletion of ovarian follicles reduces the circulating levels of inhibin A and B, and free a-subunit to nondetectable concentrations (Burger et al., 1998a). However, in postmenopausal women with granulosa cell tumors (Lappohn et al., 1989) and mucinous ovarian cancers (Healy et al., 1993), the serum levels of inhibin are substantially elevated. Thus, the measurement of inhibin in the serum of postmenopausal women may be used as a marker for ovarian tumors. A dual detection assay system that detects the inhibin a-subunit (inhibin aC-subunit ELISA; Robertson et al., 2002) and CA125, an ovarian tumor marker (Whitehouse and Solomon, 2003), provides a specific and sensitive detection system for the majority of ovarian cancers in postmenopausal women (Burger et al., 2001; Robertson et al., 2007). Inhibin detection in premenopausal women with ovarian cancers is restricted by the cyclic nature and higher levels of inhibin secretion (Robertson et al., 2007). The molecular weight forms of inhibin A, B, and pro-aC observed in granulosa cell tumors are similar to those previously identified in hFF and serum (Robertson et al., 2002). Although inhibin B is the major form of inhibin secreted by granulosa cell tumors (Burger et al., 2001; Petraglia et al., 1998; Robertson et al., 1999), mature inhibin A and pro-aC (25–35 kDa) and high molecular weight dimeric precursor forms have also been detected. In contrast, the mucinous and serous cancer serum samples contain a predominantly free a-subunit. High molecular weight inhibin forms were also observed in mucinous and serous cancer serum, whereas little inhibin A and B forms were identified. Recent studies have indicated that ovarian cancers exhibit a reduced responsiveness to inhibin signaling and that this is associated with a loss in expression of betaglycan. Hempel and colleagues reported a loss of betaglycan mRNA and protein in epithelial-derived ovarian cancer cell lines. Furthermore, reintroduction of betaglycan into these cells reduced cancer cell motility and invasiveness by specifically enhancing inhibin-mediated suppression of MMPs (Hempel et al., 2007). Similarly, recent experiments have shown that betaglycan mRNA expression is reduced in granulosa cell tumors and that cell lines derived from these tumors display poor responsiveness to inhibin A using in vitro bioassays (Bilandzic et al., 2009). The loss of inhibin activity promotes tumorigenesis; this is almost certainly a consequence of increased activin signaling in these tissues (Matzuk et al., 1992, 1994). Matzuk and colleagues have shown that in inhibin-deficient mice with tumors, the serum levels of activin are elevated by more than 10-fold (Matzuk et al., 1994).
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C. Circulating inhibin B forms in men Forms of inhibin B present in the plasma of adult males are similar to those observed in hFF and serum of females (Robertson et al., 1996, 1997). Male plasma also lacks the 55 kDa form of inhibin B, but does contain the mature inhibin B isoforms (30 kDa), and precursor forms of 100 and 20 kDa. The latter of these fragments is suggested to be free aC, based on its detection using antibodies specific for this region. The proportion of high molecular weight inhibin B appears to be higher in male serum (50%) than that of plasma (30%), suggesting that at least 50% of the circulating inhibin B is in precursor form (Robertson et al., 2003). The bioactivity of the various secreted forms of inhibin has been examined in some detail (Robertson et al., 1996, 1997). It is not only the mature inhibin A and B forms that exhibit in vitro bioactivities. Analyses by Robertson and colleagues have shown that some high molecular weight/ precursor inhibin forms retain biological activity in pituitary bioassays (Robertson et al., 1985, 1997; Sugino et al., 1992). It is possible that these high molecular weight forms are proteolytically cleaved to mature inhibin outside the cell, as has been described for other TGF-b ligands (Blanchet et al., 2008). It has been shown that the 58 kDa form of bovine inhibin A can be cleaved in serum to generate a <30 kDa form of inhibin (McLachlan et al., 1986). Recent studies have confirmed that the observed variation in bioactivities of the mature inhibin forms (31 and 34 kDa) can be attributed to their differential glycosylation (Makanji et al., 2007). Ongoing studies in our laboratory are assessing the contribution of posttranslational modifications in the differential production and bioactivity of inhibins.
VI. Concluding Remarks In summary, from its initial isolation in follicular fluid, we have gained considerable insight on the tissue-specific production of inhibins. Inhibin synthesis, occurring predominantly in the gonads, is switched on by gonadotrophins (FSH and LH) and other factors (such as the TGF-b ligands) binding to target receptors on the surface of granulosa cells of the ovary and Sertoli cells of the testis. Receptor binding initiates intracellular signaling cascades (such as the PKA pathway) that activate intracellular transcription factors (including cAMP binding proteins). Following gene activation, the inhibin mRNAs are processed by the Golgi and endoplasmic reticulum, resulting in production of the inhibin subunits (a, bA, and bB). The prodomains of the inhibin subunits direct the folding of inhibin monomers, followed by covalent dimerization of the inhibin a- and b-subunits. Following assembly, inhibins are proteolytically processed to yield various
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molecular weight isoforms of inhibin A and B. During synthesis, the inhibin subunits are also subjected to posttranslational modifications, including glycosylation. The cell-to-cell variation in inhibin subunit expression and ligand secretion is attributable to the diversity in the individual gene sequences and the factors that regulate synthesis. Regulated inhibin synthesis is paramount for the maintenance of homeostasis in reproductive tissues. Consequently, perturbed inhibin expression has been associated with reproductive disease states, such as ovarian cancers, and deleterious pregnancy outcomes. Understanding the precise mechanisms that govern inhibin synthesis and secretion will facilitate future diagnosis and treatment of such conditions.
REFERENCES Alexander, J. M., Bikkal, H. A., Zervas, N. T., Laws, E. R., Jr., and Klibanski, A. (1996). Tumor-specific expression and alternate splicing of messenger ribonucleic acid encoding activin/transforming growth factor-beta receptors in human pituitary adenomas. J. Clin. Endocrinol. Metab. 81, 783–790. Ambros, V. (2001). microRNAs: Tiny regulators with great potential. Cell 107, 823–826. Anderson, R. A., Cambray, N., Hartley, P. S., and McNeilly, A. S. (2002). Expression and localization of inhibin alpha, inhibin/activin betaA and betaB and the activin type II and inhibin beta-glycan receptors in the developing human testis. Reproduction 123, 779–788. Annes, J. P., Munger, J. S., and Rifkin, D. B. (2003). Making sense of latent TGFbeta activation. J. Cell Sci. 116, 217–224. Antenos, M., Stemler, M., Boime, I., and Woodruff, T. K. (2007). N-linked oligosaccharides direct the differential assembly and secretion of inhibin alpha- and betaA-subunit dimers. Mol. Endocrinol. 21, 1670–1684. Antenos, M., Zhu, J., Jetly, N. M., and Woodruff, T. K. (2008). An activin/furin regulatory loop modulates the processing and secretion of inhibin alpha- and betaB-subunit dimers in pituitary gonadotrope cells. J. Biol. Chem. 283, 33059–33068. Ardekani, A. M., Romanelli, J. C., and Mayo, K. E. (1998). Structure of the rat inhibin and activin betaA-subunit gene and regulation in an ovarian granulosa cell line. Endocrinology 139, 3271–3279. Attardi, B., and Miklos, J. (1990). Rapid stimulatory effect of activin-A on messenger RNA encoding the follicle-stimulating hormone beta-subunit in rat pituitary cell cultures. Mol. Endocrinol. 4, 721–726. Attardi, B., Vaughan, J., and Vale, W. (1992). Regulation of FSH beta messenger ribonucleic acid levels in the rat by endogenous inhibin. Endocrinology 130, 557–559. Ay, E., Kavak, Z. N., Elter, K., Gokaslan, H., and Pekin, T. (2005). Screening for preeclampsia by using maternal serum inhibin A, activin A, human chorionic gonadotropin, unconjugated estriol, and alpha-fetoprotein levels and uterine artery Doppler in the second trimester of pregnancy. Aust. NZ J. Obstet. Gynaecol. 45, 283–288. Balanathan, P., Ball, E. M., Wang, H., Harris, S. E., Shelling, A. N., and Risbridger, G. P. (2004). Epigenetic regulation of inhibin alpha-subunit gene in prostate cancer cell lines. J. Mol. Endocrinol. 32, 55–67. Barton, D. E., Yang-Feng, T. L., Mason, A. J., Seeburg, P. H., and Francke, U. (1989). Mapping of genes for inhibin subunits alpha, beta A, and beta B on human and mouse chromosomes and studies of jsd mice. Genomics 5, 91–99.
172
Kelly L. Walton et al.
Bersinger, N. A., Smarason, A. K., Muttukrishna, S., Groome, N. P., and Redman, C. W. (2003). Women with preeclampsia have increased serum levels of pregnancy-associated plasma protein A (PAPP-A), inhibin A, activin A and soluble E-selectin. Hypertens. Pregnancy 22, 45–55. Bhasin, S., Krummen, L. A., Swerdloff, R. S., Morelos, B. S., Kim, W. H., diZerega, G. S., Ling, N., Esch, F., Shimasaki, S., and Toppari, J. (1989). Stage dependent expression of inhibin alpha and beta-B subunits during the cycle of the rat seminiferous epithelium. Endocrinology 124, 987–991. Bicsak, T. A., Tucker, E. M., Cappel, S., Vaughan, J., Rivier, J., Vale, W., and Hsueh, A. J. (1986). Hormonal regulation of granulosa cell inhibin biosynthesis. Endocrinology 119, 2711–2719. Bicsak, T. A., Vale, W., Vaughan, J., Tucker, E. M., Cappel, S., and Hsueh, A. J. (1987). Hormonal regulation of inhibin production by cultured Sertoli cells. Mol. Cell. Endocrinol. 49, 211–217. Bilandzic, M., Chu, S., Farnworth, P. G., Harrison, C., Nicholls, P., Wang, Y., Escalona, R. M., Fuller, P. J., Findlay, J. K., and Stenvers, K. L. (2009). Loss of betaglycan contributes to the malignant properties of human granulosa tumor cells. Mol. Endocrinol. 23, 539–548. Blanchet, M. H., Le Good, J. A., Mesnard, D., Oorschot, V., Baflast, S., Minchiotti, G., Klumperman, J., and Constam, D. B. (2008). Cripto recruits Furin and PACE4 and controls Nodal trafficking during proteolytic maturation. EMBO J. 27, 2580–2591. Boitani, C., Stefanini, M., Fragale, A., and Morena, A. R. (1995). Activin stimulates Sertoli cell proliferation in a defined period of rat testis development. Endocrinology 136, 5438–5444. Bondestam, J., Kaivo-oja, N., Kallio, J., Groome, N., Hyden-Granskog, C., Fujii, M., Moustakas, A., Jalanko, A., ten Dijke, P., and Ritvos, O. (2002). Engagement of activin and bone morphogenetic protein signaling pathway Smad proteins in the induction of inhibin B production in ovarian granulosa cells. Mol. Cell. Endocrinol. 195, 79–88. Bottinger, E. P., Factor, V. M., Tsang, M. L., Weatherbee, J. A., Kopp, J. B., Qian, S. W., Wakefield, L. M., Roberts, A. B., Thorgeirsson, S. S., and Sporn, M. B. (1996). The recombinant proregion of transforming growth factor beta1 (latency-associated peptide) inhibits active transforming growth factor beta1 in transgenic mice. Proc. Natl. Acad. Sci. USA 93, 5877–5882. Brunner, A. M., Marquardt, H., Malacko, A. R., Lioubin, M. N., and Purchio, A. F. (1989). Site-directed mutagenesis of cysteine residues in the pro region of the transforming growth factor beta 1 precursor. Expression and characterization of mutant proteins. J. Biol. Chem. 264, 13660–13664. Brunner, A. M., Lioubin, M. N., Marquardt, H., Malacko, A. R., Wang, W. C., Shapiro, R. A., Neubauer, M., Cook, J., Madisen, L., and Purchio, A. F. (1992). Sitedirected mutagenesis of glycosylation sites in the transforming growth factor-beta 1 (TGF beta 1) and TGF beta 2 (414) precursors and of cysteine residues within mature TGF beta 1: Effects on secretion and bioactivity. Mol. Endocrinol. 6, 1691–1700. Burger, H. G., Robertson, D. M., Cahir, N., Mamers, P., Healy, D. L., Jobling, T., and Groome, N. (1996). Characterization of inhibin immunoreactivity in post-menopausal women with ovarian tumours. Clin. Endocrinol. (Oxf) 44, 413–418. Burger, H. G., Cahir, N., Robertson, D. M., Groome, N. P., Dudley, E., Green, A., and Dennerstein, L. (1998a). Serum inhibins A and B fall differentially as FSH rises in perimenopausal women. Clin. Endocrinol. (Oxf) 48, 809–813. Burger, H. G., Groome, N. P., and Robertson, D. M. (1998b). Both inhibin A and B respond to exogenous follicle-stimulating hormone in the follicular phase of the human menstrual cycle. J. Clin. Endocrinol. Metab. 83, 4167–4169.
Inhibin Synthesis and Secretion
173
Burger, H. G., Fuller, P. J., Chu, S., Mamers, P., Drummond, A., Susil, B., Neva, P., and Robertson, D. M. (2001). The inhibins and ovarian cancer. Mol. Cell. Endocrinol. 180, 145–148. Burkart, A. D., Mukherjee, A., Sterneck, E., Johnson, P. F., and Mayo, K. E. (2005). Repression of the inhibin alpha-subunit gene by the transcription factor CCAAT/ enhancer-binding protein-beta. Endocrinology 146, 1909–1921. Buzzard, J. J., Loveland, K. L., O’Bryan, M. K., O’Connor, A. E., Bakker, M., Hayashi, T., Wreford, N. G., Morrison, J. R., and de Kretser, D. M. (2004). Changes in circulating and testicular levels of inhibin A and B and activin A during postnatal development in the rat. Endocrinology 145, 3532–3541. Carroll, R. S., Corrigan, A. Z., Gharib, S. D., Vale, W., and Chin, W. W. (1989). Inhibin, activin, and follistatin: Regulation of follicle-stimulating hormone messenger ribonucleic acid levels. Mol. Endocrinol. 3, 1969–1976. Casagrandi, D., Bearfield, C., Geary, J., Redman, C. W., and Muttukrishna, S. (2003). Inhibin, activin, follistatin, activin receptors and beta-glycan gene expression in the placental tissue of patients with pre-eclampsia. Mol. Hum. Reprod. 9, 199–203. Centrella, M., Horowitz, M. C., Wozney, J. M., and McCarthy, T. L. (1994). Transforming growth factor-beta gene family members and bone. Endocr. Rev. 15, 27–39. Chen, C. C., and Johnson, P. A. (1996). Expression of inhibin alpha and inhibin/activin beta A subunits in the granulosa layer of the large preovulatory follicles of the hen. Biol. Reprod. 55, 450–454. Ciarmela, P., Florio, P., Toti, P., Franchini, A., Maguer-Satta, V., Ginanneschi, C., Ottaviani, E., and Petraglia, F. (2003). Human placenta and fetal membranes express follistatin-related gene mRNA and protein. J. Endocrinol. Invest. 26, 641–645. Corrigan, A. Z., Bilezikjian, L. M., Carroll, R. S., Bald, L. N., Schmelzer, C. H., Fendly, B. M., Mason, A. J., Chin, W. W., Schwall, R. H., and Vale, W. (1991). Evidence for an autocrine role of activin B within rat anterior pituitary cultures. Endocrinology 128, 1682–1684. Coss, D., Thackray, V. G., Deng, C. X., and Mellon, P. L. (2005). Activin regulates luteinizing hormone beta-subunit gene expression through Smad-binding and homeobox elements. Mol. Endocrinol. 19, 2610–2623. De Crescenzo, G., Grothe, S., Zwaagstra, J., Tsang, M., and O’Connor-McCourt, M. D. (2001). Real-time monitoring of the interactions of transforming growth factor-beta (TGF-beta ) isoforms with latency-associated protein and the ectodomains of the TGFbeta type II and III receptors reveals different kinetic models and stoichiometries of binding. J. Biol. Chem. 276, 29632–29643. Dennler, S., Itoh, S., Vivien, D., ten Dijke, P., Huet, S., and Gauthier, J. M. (1998). Direct binding of Smad3 and Smad4 to critical TGF beta-inducible elements in the promoter of human plasminogen activator inhibitor-type 1 gene. EMBO J. 17, 3091–3100. DePaolo, L. V., Bicsak, T. A., Erickson, G. F., Shimasaki, S., and Ling, N. (1991). Follistatin and activin: A potential intrinsic regulatory system within diverse tissues. Proc. Soc. Exp. Biol. Med. 198, 500–512. Derynck, R., Zhang, Y., and Feng, X. H. (1998). Smads: Transcriptional activators of TGFbeta responses. Cell 95, 737–740. Di Loreto, C., Reis, F. M., Cataldi, P., Zuiani, C., Luisi, S., Beltrami, C. A., and Petraglia, F. (1999). Human mammary gland and breast carcinoma contain immunoreactive inhibin/activin subunits: Evidence for a secretion into cystic fluid. Eur. J. Endocrinol. 141, 190–194. Drummond, A. E., Le, M. T., Ethier, J. F., Dyson, M., and Findlay, J. K. (2002). Expression and localization of activin receptors, Smads, and beta glycan to the postnatal rat ovary. Endocrinology 143, 1423–1433.
174
Kelly L. Walton et al.
Dykema, J. C., and Mayo, K. E. (1994). Two messenger ribonucleic acids encoding the common beta B-chain of inhibin and activin have distinct 50 -initiation sites and are differentially regulated in rat granulosa cells. Endocrinology 135, 702–711. Edwards, S. J., Reader, K. L., Lun, S., Western, A., Lawrence, S., McNatty, K. P., and Juengel, J. L. (2008). The cooperative effect of growth and differentiation factor-9 and bone morphogenetic protein (BMP)-15 on granulosa cell function is modulated primarily through BMP receptor II. Endocrinology 149, 1026–1030. Eramaa, M., and Ritvos, O. (1996). Transforming growth factor-beta 1 and -beta 2 induce inhibin and activin beta B-subunit messenger ribonucleic acid levels in cultured human granulosa-luteal cells. Fertil. Steril. 65, 954–960. Eramaa, M., Hilden, K., Tuuri, T., and Ritvos, O. (1995). Regulation of inhibin/activin subunit messenger ribonucleic acids (mRNAs) by activin A and expression of activin receptor mRNAs in cultured human granulosa-luteal cells. Endocrinology 136, 4382–4389. Feng, Z. M., Li, Y. P., and Chen, C. L. (1989). Analysis of the 5’-flanking regions of rat inhibin alpha- and beta-B-subunit genes suggests two different regulatory mechanisms. Mol. Endocrinol. 3, 1914–1925. Feng, Z. M., Wu, A. Z., and Chen, C. L. (1995). Characterization and regulation of two testicular inhibin/activin beta B-subunit messenger ribonucleic acids that are transcribed from alternate initiation sites. Endocrinology 136, 947–955. Florio, P., Luisi, S., D’Antona, D., Severi, F. M., Rago, G., and Petraglia, F. (2004). Maternal serum inhibin A levels may predict pregnancy outcome in women with threatened abortion. Fertil. Steril. 81, 468–470. Gleizes, P. E., Beavis, R. C., Mazzieri, R., Shen, B., and Rifkin, D. B. (1996). Identification and characterization of an eight-cysteine repeat of the latent transforming growth factorbeta binding protein-1 that mediates bonding to the latent transforming growth factorbeta1. J. Biol. Chem. 271, 29891–29896. Good, T. E., Weber, P. S., Ireland, J. L., Pulaski, J., Padmanabhan, V., Schneyer, A. L., Lambert-Messerlian, G., Ghosh, B. R., Miller, W. L., Groome, N., et al. (1995). Isolation of nine different biologically and immunologically active molecular variants of bovine follicular inhibin. Biol. Reprod. 53, 1478–1488. Gray, A. M., and Mason, A. J. (1990). Requirement for activin A and transforming growth factor–beta 1 pro-regions in homodimer assembly. Science 247, 1328–1330. Groome, N. (1991). Ultrasensitive two-site assays for inhibin-A and activin-A using monoclonal antibodies raised to synthetic peptides. J. Immunol. Methods 145, 65–69. Groome, N., and Lawrence, M. (1991). Preparation of monoclonal antibodies to the beta A subunit of ovarian inhibin using a synthetic peptide immunogen. Hybridoma 10, 309–316. Groome, N., Hancock, J., Betteridge, A., Lawrence, M., and Craven, R. (1990). Monoclonal and polyclonal antibodies reactive with the 1-32 amino terminal sequence of the alpha subunit of human 32K inhibin. Hybridoma 9, 31–42. Groome, N. P., Illingworth, P. J., O’Brien, M., Pai, R., Rodger, F. E., Mather, J. P., and McNeilly, A. S. (1996). Measurement of dimeric inhibin B throughout the human menstrual cycle. J. Clin. Endocrinol. Metab. 81, 1401–1405. Grootenhuis, A. J., Timmerman, M. A., Hordijk, P. L., and de Jong, F. H. (1990a). Inhibin in immature rat Sertoli cell conditioned medium: A 32 kDa alpha beta-B dimer. Mol. Cell. Endocrinol. 70, 109–116. Grootenhuis, A. J., van Beurden, W. M., Timmerman, M. A., and de Jong, F. H. (1990b). Follicle-stimulating hormone-stimulated secretion of an immunoreactive 29 kDa inhibin alpha-subunit complex, but not of 32 kDa bioactive inhibin, from cultured immature rat Sertoli cells. Mol. Cell. Endocrinol. 74, 125–132.
Inhibin Synthesis and Secretion
175
Groppe, J., Hinck, C. S., Samavarchi-Tehrani, P., Zubieta, C., Schuermann, J. P., Taylor, A. B., Schwarz, P. M., Wrana, J. L., and Hinck, A. P. (2008). Cooperative assembly of TGF-beta superfamily signaling complexes is mediated by two disparate mechanisms and distinct modes of receptor binding. Mol. Cell 29, 157–168. Haddad, G., Penabad, J. L., Bashey, H. M., Asa, S. L., Gennarelli, T. A., Cirullo, R., and Snyder, P. J. (1994). Expression of activin/inhibin subunit messenger ribonucleic acids by gonadotroph adenomas. J. Clin. Endocrinol. Metab. 79, 1399–1403. Hancock, A. D., Robertson, D. M., and de Kretser, D. M. (1992). Inhibin and inhibin alpha-chain precursors are produced by immature rat Sertoli cells in culture. Biol. Reprod. 46, 155–161. Hart, P. J., Deep, S., Taylor, A. B., Shu, Z., Hinck, C. S., and Hinck, A. P. (2002). Crystal structure of the human TbetaR2 ectodomain–TGF-beta3 complex. Nat. Struct. Biol. 9, 203–208. He, W. W., Gustafson, M. L., Hirobe, S., and Donahoe, P. K. (1993). Developmental expression of four novel serine/threonine kinase receptors homologous to the activin/ transforming growth factor-beta type II receptor family. Dev. Dyn. 196, 133–142. Healy, D. L., Burger, H. G., Mamers, P., Jobling, T., Bangah, M., Quinn, M., Grant, P., Day, A. J., Rome, R., and Campbell, J. J. (1993). Elevated serum inhibin concentrations in postmenopausal women with ovarian tumors. N. Engl. J. Med. 329, 1539–1542. Hempel, N., How, T., Dong, M., Murphy, S. K., Fields, T. A., and Blobe, G. C. (2007). Loss of betaglycan expression in ovarian cancer: Role in motility and invasion. Cancer Res. 67, 5231–5238. Husken-Hindi, P., Tsuchida, K., Park, M., Corrigan, A. Z., Vaughan, J. M., Vale, W. W., and Fischer, W. H. (1994). Monomeric activin A retains high receptor binding affinity but exhibits low biological activity. J. Biol. Chem. 269, 19380–19384. Illingworth, P. J., Groome, N. P., Byrd, W., Rainey, W. E., McNeilly, A. S., Mather, J. P., and Bremner, W. J. (1996). Inhibin-B: A likely candidate for the physiologically important form of inhibin in men. J. Clin. Endocrinol. Metab. 81, 1321–1325. Ireland, J. L., Good, T. E., Knight, P. G., and Ireland, J. J. (1994). Alterations in amounts of different forms of inhibin during follicular atresia. Biol. Reprod. 50, 1265–1276. Ito, M., Park, Y., Weck, J., Mayo, K. E., and Jameson, J. L. (2000). Synergistic activation of the inhibin alpha-promoter by steroidogenic factor-1 and cyclic adenosine 30 , 50 -monophosphate. Mol. Endocrinol. 14, 66–81. Jaatinen, R., Bondestam, J., Raivio, T., Hilden, K., Dunkel, L., Groome, N., and Ritvos, O. (2002). Activation of the bone morphogenetic protein signaling pathway induces inhibin beta(B)-subunit mRNA and secreted inhibin B levels in cultured human granulosa-luteal cells. J. Clin. Endocrinol. Metab. 87, 1254–1261. Jeruss, J. S., Santiago, J. Y., and Woodruff, T. K. (2003). Localization of activin and inhibin subunits, receptors and SMADs in the mouse mammary gland. Mol. Cell. Endocrinol. 203, 185–196. Jones, R. L., Salamonsen, L. A., Critchley, H. O., Rogers, P. A., Affandi, B., and Findlay, J. K. (2000). Inhibin and activin subunits are differentially expressed in endometrial cells and leukocytes during the menstrual cycle, in early pregnancy and in women using progestin-only contraception. Mol. Hum. Reprod. 6, 1107–1117. Jones, R. L., Salamonsen, L. A., and Findlay, J. K. (2002a). Potential roles for endometrial inhibins, activins and follistatin during human embryo implantation and early pregnancy. Trends Endocrinol. Metab. 13, 144–150. Jones, R. L., Salamonsen, L. A., Zhao, Y. C., Ethier, J. F., Drummond, A. E., and Findlay, J. K. (2002b). Expression of activin receptors, follistatin and betaglycan by human endometrial stromal cells; consistent with a role for activins during decidualization. Mol. Hum. Reprod. 8, 363–374.
176
Kelly L. Walton et al.
Jones, R. L., Findlay, J. K., Farnworth, P. G., Robertson, D. M., Wallace, E., and Salamonsen, L. A. (2006). Activin A and inhibin A differentially regulate human uterine matrix metalloproteinases: Potential interactions during decidualization and trophoblast invasion. Endocrinology 147, 724–732. Kaipia, A., Parvinen, M., Shimasaki, S., Ling, N., and Toppari, J. (1991). Stage-specific cellular regulation of inhibin alpha-subunit mRNA expression in the rat seminiferous epithelium. Mol. Cell. Endocrinol. 82, 165–173. Kaiser, U. B., Lee, B. L., Carroll, R. S., Unabia, G., Chin, W. W., and Childs, G. V. (1992). Follistatin gene expression in the pituitary: Localization in gonadotropes and folliculostellate cells in diestrous rats. Endocrinology 130, 3048–3056. Kaivo-Oja, N., Bondestam, J., Kamarainen, M., Koskimies, J., Vitt, U., Cranfield, M., Vuojolainen, K., Kallio, J. P., Olkkonen, V. M., Hayashi, M., Moustakas, A., Groome, N. P., et al. (2003). Growth differentiation factor-9 induces Smad2 activation and inhibin B production in cultured human granulosa-luteal cells. J. Clin. Endocrinol. Metab. 88, 755–762. Kingsley, D. M. (1994). The TGF-beta superfamily: New members, new receptors, and new genetic tests of function in different organisms. Genes Dev. 8, 133–146. Klein, N. A., Illingworth, P. J., Groome, N. P., McNeilly, A. S., Battaglia, D. E., and Soules, M. R. (1996). Decreased inhibin B secretion is associated with the monotropic FSH rise in older, ovulatory women: A study of serum and follicular fluid levels of dimeric inhibin A and B in spontaneous menstrual cycles. J. Clin. Endocrinol. Metab. 81, 2742–2745. Knight, P. G., and Glister, C. (2006). TGF-beta superfamily members and ovarian follicle development. Reproduction 132, 191–206. Knight, P. G., Beard, A. J., Wrathall, J. H., and Castillo, R. J. (1989). Evidence that the bovine ovary secretes large amounts of monomeric inhibin alpha subunit and its isolation from bovine follicular fluid. J. Mol. Endocrinol. 2, 189–200. La Rosa, S., Uccella, S., Billo, P., Facco, C., Sessa, F., and Capella, C. (1999). Immunohistochemical localization of alpha- and betaA-subunits of inhibin/activin in human normal endocrine cells and related tumors of the digestive system. Virchows Arch. 434, 29–36. La Rosa, S., Uccella, S., Marchet, S., Capella, C., and Lloyd, R. V. (2004). Localization of inhibins and activins in normal endocrine cells and endocrine tumors of the gut and pancreas: An immunohistochemical and in situ hybridization study. J. Histochem. Cytochem. 52, 217–225. Lappohn, R. E., Burger, H. G., Bouma, J., Bangah, M., Krans, M., and de Bruijn, H. W. (1989). Inhibin as a marker for granulosa-cell tumors. N. Engl. J. Med. 321, 790–793. Leung, P. H., Salamonsen, L. A., and Findlay, J. K. (1998). Immunolocalization of inhibin and activin subunits in human endometrium across the menstrual cycle. Hum. Reprod. 13, 3469–3477. Leversha, L. J., Robertson, D. M., de Vos, F. L., Morgan, F. J., Hearn, M. T., Wettenhall, R. E., Findlay, J. K., Burger, H. G., and de Kretser, D. M. (1987). Isolation of inhibin from ovine follicular fluid. J. Endocrinol. 113, 213–221. Lewis, K. A., Gray, P. C., Blount, A. L., MacConell, L. A., Wiater, E., Bilezikjian, L. M., and Vale, W. (2000). Betaglycan binds inhibin and can mediate functional antagonism of activin signalling. Nature 404, 411–414. Ling, N., Ying, S. Y., Ueno, N., Esch, F., Denoroy, L., and Guillemin, R. (1985). Isolation and partial characterization of a Mr 32, 000 protein with inhibin activity from porcine follicular fluid. Proc. Natl. Acad. Sci. USA 82, 7217–7221. Ludlow, H., Muttukrishna, S., Hyvonen, M., and Groome, N. P. (2008). Development of a new antibody to the human inhibin/activin betaB subunit and its application to improved inhibin B ELISAs. J. Immunol. Methods 329, 102–111.
Inhibin Synthesis and Secretion
177
Ludlow, H., Phillips, D. J., Myers, M., McLachlan, R. I., de Kretser, D. M., Allan, C. A., Anderson, R. A., Groome, N. P., Hyvonen, M., Colin Duncan, W., and Muttukrishna, S. (2009). A new ’total’ activin B enzyme-linked immunosorbent assay (ELISA): Development and validation for human samples. Clin. Endocrinol. (Oxf) 71, 867–873. MacConell, L. A., Leal, A. M., and Vale, W. W. (2002). The distribution of betaglycan protein and mRNA in rat brain, pituitary, and gonads: Implications for a role for betaglycan in inhibin-mediated reproductive functions. Endocrinology 143, 1066–1075. Madazli, R., Kuseyrioglu, B., Uzun, H., Uludag, S., and Ocak, V. (2005). Prediction of preeclampsia with maternal mid-trimester placental growth factor, activin A, fibronectin and uterine artery Doppler velocimetry. Int. J. Gynaecol. Obstet. 89, 251–257. Majdic, G., McNeilly, A. S., Sharpe, R. M., Evans, L. R., Groome, N. P., and Saunders, P. T. (1997). Testicular expression of inhibin and activin subunits and follistatin in the rat and human fetus and neonate and during postnatal development in the rat. Endocrinology 138, 2136–2147. Majumdar, S. S., Winters, S. J., and Plant, T. M. (1997). A study of the relative roles of follicle-stimulating hormone and luteinizing hormone in the regulation of testicular inhibin secretion in the rhesus monkey (Macaca mulatta). Endocrinology 138, 1363–1373. Makanji, Y., Harrison, C. A., Stanton, P. G., Krishna, R., and Robertson, D. M. (2007). Inhibin A and B in vitro bioactivities are modified by their degree of glycosylation and their affinities to betaglycan. Endocrinology 148, 2309–2316. Malone, F. D., Canick, J. A., Ball, R. H., Nyberg, D. A., Comstock, C. H., Bukowski, R., Berkowitz, R. L., Gross, S. J., Dugoff, L., Craigo, S. D., Timor-Tritsch, I. E., Carr, S. R., et al. (2005). First-trimester or second-trimester screening, or both, for Down’s syndrome. N. Engl. J. Med. 353, 2001–2011. Marchetti, C., Hamdane, M., Mitchell, V., Mayo, K., Devisme, L., Rigot, J. M., Beauvillain, J. C., Hermand, E., and Defossez, A. (2003). Immunolocalization of inhibin and activin alpha and betaB subunits and expression of corresponding messenger RNAs in the human adult testis. Biol. Reprod. 68, 230–235. Martins da Silva, S. J., Bayne, R. A., Cambray, N., Hartley, P. S., McNeilly, A. S., and Anderson, R. A. (2004). Expression of activin subunits and receptors in the developing human ovary: Activin A promotes germ cell survival and proliferation before primordial follicle formation. Dev. Biol. 266, 334–345. Mason, A. J., Hayflick, J. S., Ling, N., Esch, F., Ueno, N., Ying, S. Y., Guillemin, R., Niall, H., and Seeburg, P. H. (1985). Complementary DNA sequences of ovarian follicular fluid inhibin show precursor structure and homology with transforming growth factor-beta. Nature 318, 659–663. Mason, A. J., Niall, H. D., and Seeburg, P. H. (1986). Structure of two human ovarian inhibins. Biochem. Biophys. Res. Commun. 135, 957–964. Mason, A. J., Berkemeier, L. M., Schmelzer, C. H., and Schwall, R. H. (1989). Activin B: Precursor sequences, genomic structure and in vitro activities. Mol. Endocrinol. 3, 1352–1358. Mason, A. J., Farnworth, P. G., and Sullivan, J. (1996). Characterization and determination of the biological activities of noncleavable high molecular weight forms of inhibin A and activin A. Mol. Endocrinol. 10, 1055–1065. Massague, J., Cheifetz, S., Boyd, F. T., and Andres, J. L. (1990). TGF-beta receptors and TGF-beta binding proteoglycans: Recent progress in identifying their functional properties. Ann. NY Acad. Sci. 593, 59–72. Mather, J. P., Attie, K. M., Woodruff, T. K., Rice, G. C., and Phillips, D. M. (1990). Activin stimulates spermatogonial proliferation in germ-Sertoli cell cocultures from immature rat testis. Endocrinology 127, 3206–3214.
178
Kelly L. Walton et al.
Mathews, L. S., and Vale, W. W. (1991). Expression cloning of an activin receptor, a predicted transmembrane serine kinase. Cell 65, 973–982. Matsuzaki, K., Xu, J., Wang, F., McKeehan, W. L., Krummen, L., and Kan, M. (1993). A widely expressed transmembrane serine/threonine kinase that does not bind activin, inhibin, transforming growth factor beta, or bone morphogenic factor. J. Biol. Chem. 268, 12719–12723. Matzuk, M. M., Finegold, M. J., Su, J. G., Hsueh, A. J., and Bradley, A. (1992). Alphainhibin is a tumour-suppressor gene with gonadal specificity in mice. Nature 360, 313–319. Matzuk, M. M., Finegold, M. J., Mather, J. P., Krummen, L., Lu, H., and Bradley, A. (1994). Development of cancer cachexia-like syndrome and adrenal tumors in inhibindeficient mice. Proc. Natl. Acad. Sci. USA 91, 8817–8821. McLachlan, R. I., Robertson, D. M., Burger, H. G., and de Kretser, D. M. (1986). The radioimmunoassay of bovine and human follicular fluid and serum inhibin. Mol. Cell. Endocrinol. 46, 175–185. McMahon, H. E., Sharma, S., and Shimasaki, S. (2008). Phosphorylation of bone morphogenetic protein-15 and growth and differentiation factor-9 plays a critical role in determining agonistic or antagonistic functions. Endocrinology 149, 812–817. Mellor, S. L., Richards, M. G., Pedersen, J. S., Robertson, D. M., and Risbridger, G. P. (1998). Loss of the expression and localization of inhibin alpha-subunit in high grade prostate cancer. J. Clin. Endocrinol. Metab. 83, 969–975. Meunier, H., Rivier, C., Evans, R. M., and Vale, W. (1988). Gonadal and extragonadal expression of inhibin alpha, beta A, and beta B subunits in various tissues predicts diverse functions. Proc. Natl. Acad. Sci. USA 85, 247–251. Miyamoto, K., Hasegawa, Y., Fukuda, M., Nomura, M., Igarashi, M., Kangawa, K., and Matsuo, H. (1985). Isolation of porcine follicular fluid inhibin of 32K daltons. Biochem. Biophys. Res. Commun. 129, 396–403. Miyamoto, K., Hasegawa, Y., Fukuda, M., and Igarashi, M. (1986). Demonstration of high molecular weight forms of inhibin in bovine follicular fluid (bFF) by using monoclonal antibodies to bFF 32K inhibin. Biochem. Biophys. Res. Commun. 136, 1103–1109. Miyazono, K., Hellman, U., Wernstedt, C., and Heldin, C. H. (1988). Latent high molecular weight complex of transforming growth factor beta 1. Purification from human platelets and structural characterization. J. Biol. Chem. 263, 6407–6415. Molloy, S. S., Anderson, E. D., Jean, F., and Thomas, G. (1999). Bi-cycling the furin pathway: From TGN localization to pathogen activation and embryogenesis. Trends Cell Biol. 9, 28–35. Moore, K. H., Dunbar, B. S., Bousfield, G. R., and Ward, D. N. (1994). Initial characterization of equine inhibin. Biol. Reprod. 51, 63–71. Mukherjee, A., Park-Sarge, O. K., and Mayo, K. E. (1996). Gonadotropins induce rapid phosphorylation of the 3’, 5’-cyclic adenosine monophosphate response element binding protein in ovarian granulosa cells. Endocrinology 137, 3234–3245. Mukherjee, A., Urban, J., Sassone-Corsi, P., and Mayo, K. E. (1998). Gonadotropins regulate inducible cyclic adenosine 30 , 50 -monophosphate early repressor in the rat ovary: Implications for inhibin alpha subunit gene expression. Mol. Endocrinol. 12, 785–800. Muttukrishna, S., George, L., Fowler, P. A., Groome, N. P., and Knight, P. G. (1995). Measurement of serum concentrations of inhibin-A (alpha-beta A dimer) during human pregnancy. Clin. Endocrinol. (Oxf) 42, 391–397. Muttukrishna, S., Bearfield, C., Johns, J., and Jauniaux, E. (2004). Inhibin, activin, follistatin, activin receptors and beta-glycan gene expression in the villous tissue of miscarriage patients. Mol. Hum. Reprod. 10, 793–798.
Inhibin Synthesis and Secretion
179
Nakayama, K. (1997). Furin: A mammalian subtilisin/Kex2p-like endoprotease involved in processing of a wide variety of precursor proteins. Biochem. J. 327(Pt 3), 625–635. Noguchi, J., Hikono, H., Sato, S., Watanabe, G., Taya, K., Sasamoto, S., and Hasegawa, Y. (1997). Ontogeny of inhibin secretion in the rat testis: Secretion of inhibin-related proteins from fetal Leydig cells and of bioactive inhibin from Sertoli cells. J. Endocrinol. 155, 27–34. Ohga, E., Matsuse, T., Teramoto, S., and Ouchi, Y. (2000). Activin receptors are expressed on human lung fibroblast and activin A facilitates fibroblast-mediated collagen gel contraction. Life Sci. 66, 1603–1613. Okuma, Y., O’Connor, A. E., Hayashi, T., Loveland, K. L., de Kretser, D. M., and Hedger, M. P. (2006). Regulated production of activin A and inhibin B throughout the cycle of the seminiferous epithelium in the rat. J. Endocrinol. 190, 331–340. Pangas, S. A., Rademaker, A. W., Fishman, D. A., and Woodruff, T. K. (2002). Localization of the activin signal transduction components in normal human ovarian follicles: Implications for autocrine and paracrine signaling in the ovary. J. Clin. Endocrinol. Metab. 87, 2644–2657. Pei, L., Dodson, R., Schoderbek, W. E., Maurer, R. A., and Mayo, K. E. (1991). Regulation of the alpha inhibin gene by cyclic adenosine 3’, 5’-monophosphate after transfection into rat granulosa cells. Mol. Endocrinol. 5, 521–534. Penabad, J. L., Bashey, H. M., Asa, S. L., Haddad, G., Davis, K. D., Herbst, A. B., Gennarelli, T. A., Kaiser, U. B., Chin, W. W., and Snyder, P. J. (1996). Decreased follistatin gene expression in gonadotroph adenomas. J. Clin. Endocrinol. Metab. 81, 3397–3403. Petraglia, F., Vaughan, J., and Vale, W. (1989). Inhibin and activin modulate the release of gonadotropin-releasing hormone, human chorionic gonadotropin, and progesterone from cultured human placental cells. Proc. Natl. Acad. Sci. USA 86, 5114–5117. Petraglia, F., Calza, L., Garuti, G. C., Abrate, M., Giardino, L., Genazzani, A. R., Vale, W., and Meunier, H. (1990). Presence and synthesis of inhibin subunits in human decidua. J. Clin. Endocrinol. Metab. 71, 487–492. Petraglia, F., Florio, P., Luisi, S., Gallo, R., Gadducci, A., Vigano, P., Di Blasio, A. M., Genazzani, A. R., and Vale, W. (1998). Expression and secretion of inhibin and activin in normal and neoplastic uterine tissues. High levels of serum activin A in women with endometrial and cervical carcinoma. J. Clin. Endocrinol. Metab. 83, 1194–1200. Qu, J., and Thomas, K. (1995). Inhibin and activin production in human placenta. Endocr. Rev. 16, 485–507. Reame, N. E., Wyman, T. L., Phillips, D. J., de Kretser, D. M., and Padmanabhan, V. (1998). Net increase in stimulatory input resulting from a decrease in inhibin B and an increase in activin A may contribute in part to the rise in follicular phase folliclestimulating hormone of aging cycling women. J. Clin. Endocrinol. Metab. 83, 3302–3307. Reis, F. M., Cobellis, L., Tameirao, L. C., Anania, G., Luisi, S., Silva, I. S., Gioffre, W., Di Blasio, A. M., and Petraglia, F. (2002). Serum and tissue expression of activin a in postmenopausal women with breast cancer. J. Clin. Endocrinol. Metab. 87, 2277–2282. Reis, F. M., Luisi, S., Carneiro, M. M., Cobellis, L., Federico, M., Camargos, A. F., and Petraglia, F. (2004). Activin, inhibin and the human breast. Mol. Cell. Endocrinol. 225, 77–82. Rifkin, D. B. (2005). Latent transforming growth factor-beta (TGF-beta) binding proteins: Orchestrators of TGF-beta availability. J. Biol. Chem. 280, 7409–7412. Risbridger, G. P., Clements, J., Robertson, D. M., Drummond, A. E., Muir, J., Burger, H. G., and de Kretser, D. M. (1989). Immuno- and bioactive inhibin and inhibin alpha-subunit expression in rat Leydig cell cultures. Mol. Cell. Endocrinol. 66, 119–122.
180
Kelly L. Walton et al.
Rivier, J., Spiess, J., McClintock, R., Vaughan, J., and Vale, W. (1985). Purification and partial characterization of inhibin from porcine follicular fluid. Biochem. Biophys. Res. Commun. 133, 120–127. Robert, N. M., Miyamoto, Y., Taniguchi, H., and Viger, R. S. (2006). LRH-1/NR5A2 cooperates with GATA factors to regulate inhibin alpha-subunit promoter activity. Mol. Cell. Endocrinol. 257–258, 65–74. Roberts, V. J., Peto, C. A., Vale, W., and Sawchenko, P. E. (1992). Inhibin/activin subunits are costored with FSH and LH in secretory granules of the rat anterior pituitary gland. Neuroendocrinology 56, 214–224. Roberts, V. J., Barth, S., el-Roeiy, A., and Yen, S. S. (1993). Expression of inhibin/activin subunits and follistatin messenger ribonucleic acids and proteins in ovarian follicles and the corpus luteum during the human menstrual cycle. J. Clin. Endocrinol. Metab. 77, 1402–1410. Roberts, V. J., Barth, S., el-Roeiy, A., and Yen, S. S. (1994). Expression of inhibin/activin system messenger ribonucleic acids and proteins in ovarian follicles from women with polycystic ovarian syndrome. J. Clin. Endocrinol. Metab. 79, 1434–1439. Robertson, D. M., Foulds, L. M., Leversha, L., Morgan, F. J., Hearn, M. T., Burger, H. G., Wettenhall, R. E., and de Kretser, D. M. (1985). Isolation of inhibin from bovine follicular fluid. Biochem. Biophys. Res. Commun. 126, 220–226. Robertson, D. M., Giacometti, M., Foulds, L. M., Lahnstein, J., Goss, N. H., Hearn, M. T., and de Kretser, D. M. (1989). Isolation of inhibin alpha-subunit precursor proteins from bovine follicular fluid. Endocrinology 125, 2141–2149. Robertson, D., Burger, H. G., Sullivan, J., Cahir, N., Groome, N., Poncelet, E., Franchimont, P., Woodruff, T., and Mather, J. P. (1996). Biological and immunological characterization of inhibin forms in human plasma. J. Clin. Endocrinol. Metab. 81, 669–676. Robertson, D. M., Cahir, N., Findlay, J. K., Burger, H. G., and Groome, N. (1997). The biological and immunological characterization of inhibin A and B forms in human follicular fluid and plasma. J. Clin. Endocrinol. Metab. 82, 889–896. Robertson, D. M., Cahir, N., Burger, H. G., Mamers, P., and Groome, N. (1999). Inhibin forms in serum from postmenopausal women with ovarian cancers. Clin. Endocrinol. (Oxf) 50, 381–386. Robertson, D. M., Stephenson, T., Cahir, N., Tsigos, A., Pruysers, E., Stanton, P. G., Groome, N., and Thirunavukarasu, P. (2001). Development of an inhibin alpha subunit ELISA with broad specificity. Mol. Cell. Endocrinol. 180, 79–86. Robertson, D. M., Stephenson, T., Pruysers, E., McCloud, P., Tsigos, A., Groome, N., Mamers, P., and Burger, H. G. (2002). Characterization of inhibin forms and their measurement by an inhibin alpha-subunit ELISA in serum from postmenopausal women with ovarian cancer. J. Clin. Endocrinol. Metab. 87, 816–824. Robertson, D. M., Stephenson, T., and McLachlan, R. I. (2003). Characterization of plasma inhibin forms in fertile and infertile men. Hum. Reprod. 18, 1047–1054. Robertson, D. M., Pruysers, E., and Jobling, T. (2007). Inhibin as a diagnostic marker for ovarian cancer. Cancer Lett. 249, 14–17. Saharinen, J., Taipale, J., and Keski-Oja, J. (1996). Association of the small latent transforming growth factor-beta with an eight cysteine repeat of its binding protein LTBP-1. EMBO J. 15, 245–253. Saito, S., Yano, K., Sharma, S., McMahon, H. E., and Shimasaki, S. (2008). Characterization of the post-translational modification of recombinant human BMP-15 mature protein. Protein Sci. 17, 362–370. Salvador, L. M., Park, Y., Cottom, J., Maizels, E. T., Jones, J. C., Schillace, R. V., Carr, D. W., Cheung, P., Allis, C. D., Jameson, J. L., and Hunzicker-Dunn, M. (2001). Follicle-stimulating hormone stimulates protein kinase A-mediated histone H3
Inhibin Synthesis and Secretion
181
phosphorylation and acetylation leading to select gene activation in ovarian granulosa cells. J. Biol. Chem. 276, 40146–40155. Schmitt, J. F., Millar, D. S., Pedersen, J. S., Clark, S. L., Venter, D. J., Frydenberg, M., Molloy, P. L., and Risbridger, G. P. (2002). Hypermethylation of the inhibin alphasubunit gene in prostate carcinoma. Mol. Endocrinol. 16, 213–220. Sealfon, S. C., Weinstein, H., and Millar, R. P. (1997). Molecular mechanisms of ligand interaction with the gonadotropin-releasing hormone receptor. Endocr. Rev. 18, 180–205. Seder, C. W., Hartojo, W., Lin, L., Silvers, A. L., Wang, Z., Thomas, D. G., Giordano, T. J., Chen, G., Chang, A. C., Orringer, M. B., and Beer, D. G. (2009). Upregulated INHBA expression may promote cell proliferation and is associated with poor survival in lung adenocarcinoma. Neoplasia 11, 388–396. Shacham, S., Harris, D., Ben-Shlomo, H., Cohen, I., Bonfil, D., Przedecki, F., Lewy, H., Ashkenazi, I. E., Seger, R., and Naor, Z. (2001). Mechanism of GnRH receptor signaling on gonadotropin release and gene expression in pituitary gonadotrophs. Vitam. Horm. 63, 63–90. Shi, J., Yoshino, O., Osuga, Y., Koga, K., Hirota, Y., Hirata, T., Yano, T., Nishii, O., and Taketani, Y. (2009). Bone morphogenetic protein-6 stimulates gene expression of follicle-stimulating hormone receptor, inhibin/activin beta subunits, and anti-Mullerian hormone in human granulosa cells. Fertil. Steril. 92, 1794–1798. Sidis, Y., Fujiwara, T., Leykin, L., Isaacson, K., Toth, T., and Schneyer, A. L. (1998). Characterization of inhibin/activin subunit, activin receptor, and follistatin messenger ribonucleic acid in human and mouse oocytes: Evidence for activin’s paracrine signaling from granulosa cells to oocytes. Biol. Reprod. 59, 807–812. Spencer, S. J., Rabinovici, J., and Jaffe, R. B. (1990). Human recombinant activin-A inhibits proliferation of human fetal adrenal cells in vitro. J. Clin. Endocrinol. Metab. 71, 1678–1680. Spencer, S. J., Rabinovici, J., Mesiano, S., Goldsmith, P. C., and Jaffe, R. B. (1992). Activin and inhibin in the human adrenal gland. Regulation and differential effects in fetal and adult cells. J. Clin. Invest. 90, 142–149. Spencer, K., Yu, C. K., Savvidou, M., Papageorghiou, A. T., and Nicolaides, K. H. (2006). Prediction of pre-eclampsia by uterine artery Doppler ultrasonography and maternal serum pregnancy-associated plasma protein-A, free beta-human chorionic gonadotropin, activin A and inhibin A at 22 þ 0 to 24 þ 6 weeks’ gestation. Ultrasound Obstet. Gynecol. 27, 658–663. Steinberger, A. (1979). Inhibin production by Sertoli cells in culture. J. Reprod. Fertil. Suppl. 26, 31–45. Stewart, A. G., Milborrow, H. M., Ring, J. M., Crowther, C. E., and Forage, R. G. (1986). Human inhibin genes. Genomic characterisation and sequencing. FEBS Lett. 206, 329–334. Sugino, K., Nakamura, T., Takio, K., Miyamoto, K., Hasegawa, Y., Igarashi, M., Titani, K., and Sugino, H. (1992). Purification and characterization of high molecular weight forms of inhibin from bovine follicular fluid. Endocrinology 130, 789–796. Suzuki, T., Miyamoto, K., Hasegawa, Y., Abe, Y., Ui, M., Ibuki, Y., and Igarashi, M. (1987). Regulation of inhibin production by rat granulosa cells. Mol. Cell. Endocrinol. 54, 185–195. Tanimoto, K., Handa, S., Ueno, N., Murakami, K., and Fukamizu, A. (1991). Structure and sequence analysis of the human activin beta A subunit gene. DNA Seq. 2, 103–110. Tanimoto, K., Yoshida, E., Mita, S., Nibu, Y., Murakami, K., and Fukamizu, A. (1996). Human activin betaA gene. Identification of novel 5’ exon, functional promoter, and enhancers. J. Biol. Chem. 271, 32760–32769.
182
Kelly L. Walton et al.
Thirunavukarasu, P., Stephenson, T., Forray, J., Stanton, P. G., Groome, N., Wallace, E., and Robertson, D. M. (2001). Changes in molecular weight forms of inhibin A and proalpha C in maternal serum during human pregnancy. J. Clin. Endocrinol. Metab. 86, 5794–5804. Thirunavukarasu, P., Robertson, D. M., Dole, A., Waldron, K., Dawson, G., and Wallace, E. M. (2003). Placental, maternal serum and amniotic fluid molecular weight forms of inhibin A and pro-alphaC. Placenta 24, 370–377. Thomas, T. Z., Chapman, S. M., Hong, W., Gurusingfhe, C., Mellor, S. L., Fletcher, R., Pedersen, J., and Risbridger, G. P. (1998). Inhibins, activins, and follistatins: Expression of mRNAs and cellular localization in tissues from men with benign prostatic hyperplasia. Prostate 34, 34–43. Tibaldi, E., Arrigoni, G., Martinez, H. M., Inagaki, K., Shimasaki, S., and Pinna, L. A. (2010). Golgi apparatus casein kinase phosphorylates bioactive Ser-6 of bone morphogenetic protein 15 and growth and differentiation factor 9. FEBS Lett. 584(4), 801–805. Tremblay, J. J., and Viger, R. S. (2001). GATA factors differentially activate multiple gonadal promoters through conserved GATA regulatory elements. Endocrinology 142, 977–986. Tsonis, C. G., Hillier, S. G., and Baird, D. T. (1987). Production of inhibin bioactivity by human granulosa-lutein cells: Stimulation by LH and testosterone in vitro. J. Endocrinol. 112, R11–R14. Tuuri, T., Eramaa, M., Hilden, K., and Ritvos, O. (1994). The tissue distribution of activin beta A- and beta B-subunit and follistatin messenger ribonucleic acids suggests multiple sites of action for the activin-follistatin system during human development. J. Clin. Endocrinol. Metab. 78, 1521–1524. Uccella, S., La Rosa, S., Genasetti, A., and Capella, C. (2000). Localization of inhibin/ activin subunits in normal pituitary and in pituitary adenomas. Pituitary 3, 131–139. van Schaik, R. H., Wierikx, C. D., Timmerman, M. A., Oomen, M. H., van Weerden, W. M., van der Kwast, T. H., van Steenbrugge, G. J., and de Jong, F. H. (2000). Variations in activin receptor, inhibin/activin subunit and follistatin mRNAs in human prostate tumour tissues. Br. J. Cancer 82, 112–117. Vliegen, M. K., Schlatt, S., Weinbauer, G. F., Bergmann, M., Groome, N. P., and Nieschlag, E. (1993). Localization of inhibin/activin subunits in the testis of adult nonhuman primates and men. Cell Tissue Res. 273, 261–268. Wada, M., Shintani, Y., Kosaka, M., Sano, T., Hizawa, K., and Saito, S. (1996). Immunohistochemical localization of activin A and follistatin in human tissues. Endocr. J. 43, 375–385. Wakefield, L. M., Smith, D. M., Flanders, K. C., and Sporn, M. B. (1988). Latent transforming growth factor-beta from human platelets. A high molecular weight complex containing precursor sequences. J. Biol. Chem. 263, 7646–7654. Wallace, E. M., Swanston, I. A., McNeilly, A. S., Ashby, J. P., Blundell, G., Calder, A. A., and Groome, N. P. (1996). Second trimester screening for Down’s syndrome using maternal serum dimeric inhibin A. Clin. Endocrinol. (Oxf). 44, 17–21. Wallace, E. M., Marjono, B., Tyzack, K., and Tong, S. (2004). First trimester levels of inhibins and activin A in normal and failing pregnancies. Clin. Endocrinol. (Oxf). 60, 484–490. Walsh, S., Metter, E. J., Ferrucci, L., and Roth, S. M. (2007). Activin-type II receptor B (ACVR2B) and follistatin haplotype associations with muscle mass and strength in humans. J. Appl. Physiol. 102, 2142–2148. Walton, K. L., Makanji, Y., Wilce, M. C., Chan, K. L., Robertson, D. M., and Harrison, C. A. (2009). A common biosynthetic pathway governs the dimerization and secretion of inhibin and related transforming growth factor beta (TGFbeta) ligands. J. Biol. Chem. 284, 9311–9320.
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Walton, K. L., Makanji, Y., Chen, J., Wilce, M. C., Chan, K. L., Robertson, D. M., and Harrison, C. A. (2010). Two distinct regions of latency associated peptide coordinate stability of the latent TGF-{beta}1 complex. J. Biol. Chem. 285, 17029–17037. Wang, H., and Tsang, B. K. (2007). Nodal signalling and apoptosis. Reproduction 133, 847–853. Wang, H., Jiang, J. Y., Zhu, C., Peng, C., and Tsang, B. K. (2006). Role and regulation of nodal/activin receptor-like kinase 7 signaling pathway in the control of ovarian follicular atresia. Mol. Endocrinol. 20, 2469–2482. Welt, C. K. (2004). Regulation and function of inhibins in the normal menstrual cycle. Semin. Reprod. Med. 22, 187–193. Welt, C. K., and Schneyer, A. L. (2001). Differential regulation of inhibin B and inhibin a by follicle-stimulating hormone and local growth factors in human granulosa cells from small antral follicles. J. Clin. Endocrinol. Metab. 86, 330–336. Welt, C. K., Smith, Z. A., Pauler, D. K., and Hall, J. E. (2001). Differential regulation of inhibin A and inhibin B by luteinizing hormone, follicle-stimulating hormone, and stage of follicle development. J. Clin. Endocrinol. Metab. 86, 2531–2537. Welt, C. K., Pagan, Y. L., Smith, P. C., Rado, K. B., and Hall, J. E. (2003). Control of follicle-stimulating hormone by estradiol and the inhibins: Critical role of estradiol at the hypothalamus during the luteal-follicular transition. J. Clin. Endocrinol. Metab. 88, 1766–1771. Whitehouse, C., and Solomon, E. (2003). Current status of the molecular characterization of the ovarian cancer antigen CA125 and implications for its use in clinical screening. Gynecol. Oncol. 88, S152–S157. Winters, S. J., and Plant, T. M. (1999). Partial characterization of circulating inhibin-B and pro-alphaC during development in the male rhesus monkey. Endocrinology 140, 5497–5504. Woodruff, T. K., Meunier, H., Jones, P. B., Hsueh, A. J., and Mayo, K. E. (1987). Rat inhibin: Molecular cloning of alpha- and beta-subunit complementary deoxyribonucleic acids and expression in the ovary. Mol. Endocrinol. 1, 561–568. Woodruff, T. K., Besecke, L. M., Groome, N., Draper, L. B., Schwartz, N. B., and Weiss, J. (1996). Inhibin A and inhibin B are inversely correlated to follicle-stimulating hormone, yet are discordant during the follicular phase of the rat estrous cycle, and inhibin A is expressed in a sexually dimorphic manner. Endocrinology 137, 5463–5467. Wrana, J. L., Attisano, L., Wieser, R., Ventura, F., and Massague, J. (1994). Mechanism of activation of the TGF-beta receptor. Nature 370, 341–347. Yamada, Y., Yamamoto, H., Yonehara, T., Kanasaki, H., Nakanishi, H., Miyamoto, E., and Miyazaki, K. (2004). Differential activation of the luteinizing hormone beta-subunit promoter by activin and gonadotropin-releasing hormone: A role for the mitogenactivated protein kinase signaling pathway in LbetaT2 gonadotrophs. Biol. Reprod. 70, 236–243. Yang, Z., Mu, Z., Dabovic, B., Jurukovski, V., Yu, D., Sung, J., Xiong, X., and Munger, J. S. (2007). Absence of integrin-mediated TGFbeta1 activation in vivo recapitulates the phenotype of TGFbeta1-null mice. J. Cell Biol. 176, 787–793. Ying, S. Y., Czvik, J., Becker, A., Ling, N., Ueno, N., and Guillemin, R. (1987). Secretion of follicle-stimulating hormone and production of inhibin are reciprocally related. Proc. Natl. Acad. Sci. USA 84, 4631–4635. Yndestad, A., Ueland, T., Oie, E., Florholmen, G., Halvorsen, B., Attramadal, H., Simonsen, S., Froland, S. S., Gullestad, L., Christensen, G., Damas, J. K., and Aukrust, P. (2004). Elevated levels of activin A in heart failure: Potential role in myocardial remodeling. Circulation 109, 1379–1385.
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Young, G. D., and Murphy-Ullrich, J. E. (2004). The tryptophan-rich motifs of the thrombospondin type 1 repeats bind VLAL motifs in the latent transforming growth factor-beta complex. J. Biol. Chem. 279, 47633–47642. Yu, A. W., Shao, L. E., Frigon, N. L., Jr., and Yu, J. (1994). Detection of functional and dimeric activin A in human marrow microenvironment. Implications for the modulation of erythropoiesis.. Ann. NY Acad. Sci. 718, 285–298(discussion 298–9). Zawel, L., Dai, J. L., Buckhaults, P., Zhou, S., Kinzler, K. W., Vogelstein, B., and Kern, S. E. (1998). Human Smad3 and Smad4 are sequence-specific transcription activators. Mol. Cell 1, 611–617. Zhang, Z. W., Carson, R. S., Herington, A. C., Lee, V. W., and Burger, H. G. (1987). Follicle-stimulating hormone and somatomedin-C stimulate inhibin production by rat granulosa cells in vitro. Endocrinology 120, 1633–1638.
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Multifunctional Roles of Activins in the Brain Hiroshi Ageta and Kunihiro Tsuchida Contents I. II. III. IV.
Introduction Expression Pattern of Activin and Activin Receptor in the Brain Activin Receptor and its Regulatory Proteins Functions of Activins in the CNS A. Activin regulates spine formation B. Activin influences depression and anxiety-related behavior C. Activin is an important factor in adult neurogenesis D. Activin is a key player for maintaining late-phase LTP E. Activin influences reconsolidation and extinction V. Conclusion and Perspectives Acknowledgments References
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Abstract Activins, which are members of the TGF-b superfamily, were initially isolated from gonads and served as modulators of follicle-stimulating hormone secretion. Activins regulate various biological functions, including induction of the dorsal mesoderm, craniofacial development, and differentiation of numerous cell types. Activin receptors are highly expressed in neuronal cells, and activin mRNA expression is upregulated by neuronal activity. Activins also exhibit neuroprotective action during excitotoxic brain injury. However, very little is known about the functional roles of activins in the brain. We recently generated various types of transgenic mice, demonstrating that activins regulate spine formation, behavioral activity, anxiety, adult neurogenesis, latephase long-term potentiation, and maintenance of long-term memory. The present chapter describes recent progress in the study of the role of activin in the brain. ß 2011 Elsevier Inc.
Division for Therapies against Intractable Diseases, Institute for Comprehensive Medical Science (ICMS), Fujita Health University, Toyoake, Aichi, Japan Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00009-3
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2011 Elsevier Inc. All rights reserved.
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I. Introduction In 1986, during inhibin purification, activin, a regulating factor for secretion of follicle-stimulating hormone from pituitary cells was discovered (Ling et al., 1986; Vale et al., 1986). For more than 20 years after its discovery, numerous studies have shown that activins serve as multifunctional growth and differentiation factors in many cell types (Mather et al., 1997; Ying et al., 1997). Activins are dimeric glycoproteins, which are formed by two of four different b subunits of inhibin in mammals (bA, bB, bC, and bE; Tsuchida, 2004). bA and bB transcripts exist in almost all tissues. In contrast, bC and bE subunits are predominantly expressed in the liver. Homodimers of inhibin bA or bB subunits, activin A and activin B, respectively, exist in various tissues (Nakamura et al., 1992), and heterodimeric activin AB has also been isolated from porcine follicular fluid (Ling et al., 1986; Nakamura et al., 1992). Activins directly bind to serine/threonine kinase activin type II receptors (ActRII and ActRIIB), which are located on the cell membrane (Pangas and Woodruff, 2000; Fig. 9.1). Once the ligand is bound, type II receptors
Activin A Follistatin Ca
2+
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GKAP
ActRIB/ALK4 ActR II
PSD-95
ARIP1/S-SCAM
P
P
P Smad 2/3 Smad 4
ERK F-actin organization
Transcriptional regulation
Figure 9.1 Activin-signaling cascade in the postsynaptic region. Activin A binds directly to activin type II receptor (ActRII or ActRIIB). Following binding, ActRIIs phosphorylate activin type I receptors (ActRIB or ALK4). ActRIB phosphorylates the transcriptional factor Smad2/3, and phosphorylated Smad2/3 binds to Smad4. Smad complexes translocate to the nucleus and regulate transcriptional activities. In addition, activins activate ERK signaling and lead to NMDA receptor phosphorylation in neurons. Follistatin specifically binds to activins. The activin-bound follistatin does not have access to ActRIIs. Therefore, follistatins are endogenous inhibitors of activin signals. ActRIIs form large protein complexes, including NMDAR, ARIP1, and PSD95.
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recruit and phosphorylate activin type I receptors (ActRIs), termed activin receptor-like kinase 4 (ALK4 also known as ActRIB) for activin A or ALK7 for activin B. Activated ActRIs then phosphorylate transcriptional factors Smad2/3, and phosphorylated Smad2/3 binds to Smad4. Subsequently, Smad complexes are translocated to the nucleus to regulate transcriptional activities. Follistatins, which are also secreted factors, bind to activins with high affinity. The activin-bound follistatin does not have access to activin receptors (Fig. 9.1). Therefore, follistatins are specific inhibitors for the activin-signaling cascade. Inhibin bA KO mice die within 24 h after birth, and Follistatin KO mice exhibit growth retardation and die within hours after birth, which is due to respiratory failure (Matzuk et al., 1995a,b), demonstrating that activin signals are important for normal embryonic and postnatal development. Activin signals are also involved in the pathogenesis of a variety of diseases, including metabolic diseases, musculoskeletal disorders, cancers, and mental disorders. According to recent studies, activin signaling could be a promising target for these disorders (Tsuchida et al., 2009). Recently, we generated several types of transgenic mice, which demonstrated that activins regulate spine formation, behavioral activity, anxiety, adult neurogenesis, late-phase long-term potentiation (LTP), and maintenance of long-term memory (LTM). This chapter provides an overview of recent progress in the study of the role of activin in the brain.
II. Expression Pattern of Activin and Activin Receptor in the Brain In 1995, Andreasson and Worley isolated the neural activity-dependent gene inhibin bA mRNA and showed that its regulation was dependent upon N-methyl-D-aspartate (NMDA) receptor activation (see Section III about NMDA receptor; Andreasson and Worley, 1995). At the same time using the differential display method, inhibin bA mRNA was identified as a neural activity-dependent gene in the rat hippocampus (Inokuchi et al., 1996). In contrast, however, inhibin a mRNA levels were not affected by neuronal activity (Inokuchi et al., 1996). In addition, activin receptor ActRII mRNA is expressed in the adult brain and is specifically abundant in the hippocampus and amygdala (Cameron et al., 1994). Immunohistochemical analysis also revealed ActRII expression in neurons of the cerebral cortex, hippocampus, medial amygdala, and thalamus (Funaba et al., 1997).
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III. Activin Receptor and its Regulatory Proteins The brain is composed of neurons that communicate with one another by transmitting chemicals (neurotransmitters) (Fig. 9.2A). To accomplish this, neurons develop two distinct processes—axons and dendrites (Fig. 9.2A). Information flows from one neuron to another across a synapse, which is a small gap between the neurons. Chemicals (neurotransmitters) are released from the axonal terminal (presynapse), which then bind to cell-surface receptors located on the dendrites (postsynapse; Fig. 9.2A). Spines are small, membranous protrusions on the dendrite. The majority of spines have a bulbous head (the spine head) and a thin neck that connects the head of the spine to the dendrite shaft (Fig. 9.2A). In the spine head, crucial proteins accumulate to respond to presynaptic regional signals, which are linked by scaffold proteins.
A
B
Dendrite
Pre
Post
Axon Synapse
Pre synapse
Spine
Neck
Post synapse
Activin treatment Elongation
Figure 9.2 Activins regulate spine morphology. (A) Illustration of neuron, synapse, and spine structures. (B) Cultured hippocampal neurons were treated with vehicle (upper) and activins (lower), followed by staining with phalloidin to identify F-actin (red, blue arrows) and antisynaptophysin antibody (green, white arrows) to identify presynaptic regions. Scale bar, 1 mm.
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Activin type II receptor, ActRII, binds to a scaffold protein called activin receptor interacting protein 1 (ARIP1)/synaptic scaffolding molecule (S-SCAM)/membrane-associated guanylate kinase, WW and PDZ domain containing 2 (MAGI2), which is localized at the postsynaptic site (Fig. 9.1). In addition, ARIP1 binds to guanylate kinase domain-associated protein (GKAP), Fyn kinase (Kurisaki et al., 2008), and Smad3 (Shoji et al., 2000). Postsynaptic density 95 (PSD95) is a well-characterized scaffold protein in the postsynaptic region, which binds to various protein types, including channel/receptor proteins (e.g., K channel, NMDA receptor, and Erb4), adhesion proteins (e.g., neuroligin), signaling proteins (e.g., synaptic RasGAP (SynGap) and neuronal nitric oxide synthase (nNOS)), and other neuronal function regulators (e.g., Stargazin and GKAP; Hata and Takai, 1999; Sheng and Sala, 2001; Talmage, 2008). GKAP binds to numerous proteins via Shank (Tu et al., 1999) and Vesl-1L/Homer-1C (Kato et al., 1998). Therefore, ActRII is a component of the huge PSD-95 protein complex located in the postsynaptic site. Glutamate is the major excitatory neurotransmitter in the mammalian central nerve system (CNS). It acts via two classes of receptors—ligandgated ion channels (ionotropic receptors) and G-protein-coupled metabotropic receptors. The ionotropic glutamate receptors are subdivided into three groups (AMPA, NMDA, and kainate receptors) based on pharmacological properties. NMDA receptors are highly expressed in the hippocampus, and dysregulated activation and/or inhibition of NMDA receptors influences many CNS disorders, including stroke, Parkinson’s disease, Alzheimer’s disease, epilepsy, drug dependence, depression, anxiety, and schizophrenia (Muir, 2006; Parsons et al., 1999; Sawa and Snyder, 2002; Skolnick, 1999). Activation of NMDA receptors results in influx of Ca2þ. Ca2þ influx through NMDA receptors is thought to play critical roles for memory acquisition and LTP (see Section IV.D for details). Activin treatment induces phosphorylation of NMDA receptors in primary hippocampal cultures, which is dependent on Fyn tyrosine kinase and ARIP1. In addition, activins increase Ca2þ influx through these NMDA receptor complexes (Kurisaki et al., 2008). These results indicate that activins influence neuronal activity and are involved in a multitude of CNS disorders. According to our recent work, activins also play a role in anxiety, memory, and LTP (see below).
IV. Functions of Activins in the CNS A. Activin regulates spine formation Individual spines, which are comprised of cytoskeletal actin, undergo actindependent shape changes that are regulated by neurotransmitter stimulation. This phenomenon could contribute to plasticity of brain circuits (Fischer et al., 1998, 2000; Fukazawa et al., 2003; Honkura et al., 2008; Matus, 2005).
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In typical hippocampal cultures, the majority of spine contact takes place at only one presynaptic site. Activin treatment enhances the number of presynaptic contacts per individual spine, which elevates the average length of the spine neck through F-actin organization (Fig. 9.2B; Shoji-Kasai et al., 2007). In addition to the canonical Smad pathway, activin receptors activate other Smad-independent pathways (p38 MAPK, ERK1/2, and JNK) in a cell type-specific manner (Bao et al., 2005; de Guise et al., 2006; ten Dijke et al., 2000; Werner and Alzheimer, 2006). The influence of activins on spinal morphology is independent of protein and RNA synthesis (ShojiKasai et al., 2007). However, it is interesting to note that these phenomena are completely blocked by treatment with MEK inhibitor. Following treatment with activins, ERK1/2 phosphorylation is markedly increased in hippocampal cultures, but not in astroglial-enriched cultures. However, activins do not result in a significant increase in JNK and p38 MAPK phosphorylation. These results suggested that ERK pathways primarily affect activin-dependent spine changes in hippocampal neurons.
B. Activin influences depression and anxiety-related behavior Mood disorders, such as bipolar disorder and depression, represent one of the most common mental illnesses, affecting as many as 17% individuals in the United States (Kessler et al., 1994). Because depression is a leading cause of suicide, it is considered a serious disorder in today’s society. Previous results have shown that activin A infusion into the rat hippocampus produced an antidepressant-like effect in the forced swimming test, which is typically used to assess the effects of antidepressant drugs (Dow et al., 2005). We have generated activin and follistatin transgenic mice under the control of the aCaMKII promoter, whose activity is thought to influence postnatal development in the forebrain. Several behavioral analyses were performed on these mice—namely, open field test, light and dark choice test, elevated plus maze test, and novel-area accessing test (Ageta et al., 2008). The open field test is used to measure locomotor activity, and light and dark choice test, elevated plus maze test, and novel-area accessing test are used to measure anxiety levels. Results demonstrated that follistatin overexpression mice (FSM) exhibit decreased general locomotor activity and enhanced anxiety. In contrast, activin overexpression mice (ACM) exhibit more aggressive behavior than wild-type littermates, as well as reduced anxiety-related behavior. These results showed that activin levels in the forebrain affect locomotor activity and anxiety-related behavior (Fig. 9.3 and Table 9.1). Other studies have generated dominant-negative ActRIB transgenic mice under control of the aCaMKII promoter, which demonstrated that activin signals are negative regulators of anxiety (Zheng et al., 2009). For
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FSM: forebrain specific follistatin transgenic mice ACM: forebrain specific activin transgenic mice
Figure 9.3 Activins influence anxiety-related behavior. (A) Open field test. (B) Light and dark test. Right panels show typical traces for each genotype.
example, these mice visited a greater number of inner fields in the open field test, which represents risk-taking behavior, and the mice also spent more time on lit, elevated places in the light-dark exploration test. These behaviors correlate with the level of anxiety in an animal, and results from this study were consistent with results from our laboratory. However, the conflicting results between the two studies could be due to transgene differences; Zheng et al. utilized the dominant-negative receptor ActRIB as a transgene. By contrast, we have utilized secreted factors (activins and follistatins) to determine the role of activin in the brain. GDF11, another member of the TGF-b superfamily, is expressed in the adult brain and it also binds to activin receptor II and IB. (Nakashima et al., 1999). Therefore, differences in results could be due to regulation of GDF11, as well as activins. It is possible that the dominant-negative receptor ActRIB dimerizes with receptors of the TGF-b family members other than activins. Alternatively, the dominant-negative ActRIB lacks a cytoplasmic kinase domain, thereby blocking the activin-Smad signaling cascade (see above, Fig. 9.1). Recent studies have shown that Smurf1, an E3 ubiquitin ligase, associates with the cytoplasmic domain of the TGF-b type I receptor and induces internalization and degradation of TGF-b receptors (Di Guglielmo et al., 2003; Ebisawa et al., 2001; ten Dijke and Hill, 2004). ActRIB also binds to and is ubiquitinated by the Smad7–Smurf1 complex via the ActRIB cytoplasmic domain (Yamaguchi et al., 2006). Furthermore, activin-dependent spine changes are independent of Smad activation in hippocampal cultures (Shoji-Kasai et al., 2007) (see above, Fig. 9.2). In vivo, if activin receptors are regulated by this kind of ubiquitination and proteasomal degradation,
Table 9.1
Summary of behavioral analysis of FSM and ACM
Open field test
FSM ACM
Light and dark test
Elevated plus-maze test
Novel-area accessing test
Walking speed
Time spent in locomotion
Time in rearing
Risk-taking behavior
Time in light compartment
Time in open arm
Time in novel area
– –
# –
# "
# "
# "
– "
# NT
# downregulated compared with wild-type mice; " upregulated compared with wild-type mice; – not significant change in behavior; NT, not tested; ACM, forebrainspecific activin transgenic mice; FSM, forebrain-specific follistatin transgenic mice.
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expression of dominant-negative ActRIB could lead to inhibited degradation and extension of the activation period, thereby inducing an ACM phenocopy. Previous studies have suggested correlations between anxiety and depression. Many popularly prescribed antidepressant drugs modulate monoamine neurotransmission, which could take 6–8 weeks to exert effects, and individual drugs are efficacious in only 60–70% of patients. A novel antidepressant, which acts rapidly and safely in a high proportion of patients, would be highly advantageous (Wong and Licinio, 2004). Therefore, activins and the involved signaling pathway could represent novel therapeutic targets for depression, as well as ischemic brain injury. The transgenic mice generated in our laboratory could be useful for screening compounds for newly developed and novel antidepressant drugs (Ageta et al., 2008).
C. Activin is an important factor in adult neurogenesis Postnatal neurogenesis, which is the production of new neurons in the adult brain, influences a number of physiological roles, including the replacement of damaged neurons, stress responses, and memory formation (Gage, 2002; Kitamura et al., 2009; Kokaia and Lindvall, 2003; Shors et al., 2001; Wang et al., 2004). Several recent reports have demonstrated that neurogenesis is also involved in depression (Malberg et al., 2000; Santarelli et al., 2003). Bromodeoxyuridine (BrdU), which is a thymidine analog, is incorporated into genomic DNA during S-phase and can be used as a marker of dividing cells. Through double-labeling and immunofluorescent methods utilizing antibodies specific for BrdU and the mature neuronal marker NeuN, newly generated neurons can be easily detected within brain tissue sections (Kuhn et al., 1996). In 2000, chronic antidepressant treatment was shown to significantly increase the number of BrdU-labeled cells in the hippocampal dentate gyrus and hilus (Malberg et al., 2000). An additional study demonstrated that the disruption of antidepressant-induced neurogenesis actually blocks the behavioral effects of antidepressants (Santarelli et al., 2003). Recently, our laboratory showed that activins are involved in adult neurogenesis (Ageta et al., 2008); FSM and ACM 5-week-old transgenic mice were injected with 75 mg/kg BrdU 3 times per day for 3 days. These mice were sacrificed either 24 h or 4 weeks after the final injection; at 24 h after the final injection, BrdU incorporation was measured in the hippocampal subgranular zone (SGZ), where dividing progenitor cells are located (Gage, 2002). There was no change in the number of BrdU-positive cells in the transgenic mice compared with wild-type mice at 24 h. However, after 4 weeks, the number of BrdU/NeuN-positive cells was significantly decreased in the FSM mice, but there was no neurogenesis effect in the ACM mice.
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These results indicated that endogenous activin signals are essential for adult neurogenesis (Ageta et al., 2008). In addition, the level of neurogenesis in FSM/ACM-double transgenic mice was significantly greater than in FSM mice, indicating that reduced neurogenesis in the FSM mice was partially rescued by increased activin expression (Ageta et al., 2008). Similar results have been reported by other groups (Abdipranoto-Cowley et al., 2009). These studies suggested that activin is an important factor for adult neurogenesis. Follistatin inhibits activins, but it also antagonizes other members of the TGF-b superfamily, namely, GDF11/BMP11 (Gamer et al., 1999) and GDF8/myostatin (Tsuchida, 2004). Because increased neurogenesis was not observed in the activin overexpressing mice (ACM), the possibility of decreased neurogenesis in follistatin overexpressing mice (FSM) due to GDF11 or GDF8 could not be ruled out. However, the likelihood of this is quite low, because of the following: (1) GDF8 is exclusively expressed in skeletal muscle (McPherron et al., 1997; Sharma et al., 1999); (2) GDF11 is mainly expressed in olfactory epithelium (OE) in adult mice; GDF11 mRNA expression is low in the hippocampus (ALLEN Brain project: http://www.brain-map.org); and (3) GDF11 acts as a negative regulator for neurogenesis in the OE. In fact, GDF11 inhibits production of progenitors and neurons (Wu et al., 2003), which was shown in mice lacking follistatin. In FSM mice, the number of progenitor cells is normal, but survival of newly generated neurons is significantly decreased (Ageta et al., 2008). Therefore, it is likely that activins, not GDF8 and GDF11, regulate adult hippocampal neurogenesis. Results from our study demonstrated that decreased postnatal neurogenesis, which is due to activin inhibition, results in anxiety-related behavior during adulthood (Fig. 9.8). Activin treatment in hippocampal cultures suppresses emergence of GAD67(þ) GABAergic neurons and increases the percentage of Prox1 (þ), dentate granule neurons. In contrast, follistatin treatment increases the percentage of GAD67(þ) neurons and decreases the percentage of Prox1(þ) neurons (Sekiguchi et al., 2009). These results indicated that activin signaling during postnatal neural development alters neural circuitry composition by regulating the ratio of excitatory to inhibitory neurons. In addition, results have shown that GABAergic neurotransmission is altered in dominant-negative ActRIB transgenic mice (Zheng et al., 2009). Activins increase synaptic input for each individual spine (Fig. 9.2; Shoji-Kasai et al., 2007), Ca2þ influx via the NMDA receptor (Kurisaki et al., 2008), ratio of excitatory to inhibitory neurons in newly generated neurons (Sekiguchi et al., 2009), and maintenance of early-phase LTP (E-LTP; see below) (Ageta et al., 2010). When the summation of excitatory and inhibitory postsynaptic potential from numerous synaptic inputs reaches the triggering threshold, the action
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potential propagates through the axon and triggers neurotransmitter release from the presynaptic region. Therefore, these activin-dependent actions positively affect action potential activation; in other words, activins induce activation of the entire neuronal circuit. Because electroconvulsive therapy (ECT) is used most often to treat severe major depression that does not respond to other treatments (Mukherjee et al., 1994), the resulting activin-dependent changes could positively affect depression. If this were the case, the combination of compounds that enhance activin signals and low-level electroconvulsive stimulation may be suitable for human therapy, because low-level stimulation reduces unpredictable side effects. In addition, the treatment of compounds that enhance activin signals could be utilized to replace ECT.
D. Activin is a key player for maintaining late-phase LTP LTP, which is thought to underlie learning and memory mechanisms, exhibits two distinct phases, named early-phase LTP (E-LTP) and latephase LTP (L-LTP). E-LTP persists for several hours and does not depend on protein synthesis, whereas L-LTP persists for weeks and depends on de novo RNA transcription and protein synthesis (Abraham et al., 1993; Frey et al., 1988; Nguyen et al., 1994). The formation of LTM also requires de novo RNA transcription and protein synthesis (Bourtchuladze et al., 1994; Castellucci et al., 1989; Squire and Barondes, 1973). Therefore, activitydependent gene expression is expected to play a crucial role in LTM. To understand the molecular mechanisms of LTM, many studies have isolated a number of neuronal activity-dependent genes, including inhibin bA (Andreasson and Worley, 1995; Inokuchi et al., 1996), vesl-1s/homer-1a (Brakeman et al., 1997; Kato et al., 1997), zif268 (Cole et al., 1989), arc (Lyford et al., 1995), and scrapper (Yao et al., 2007). Arc and zif268-deficient mice exhibit impaired long-term memory, but not short-term memory ( Jones et al., 2001; Plath et al., 2006). Vesl-1S protein is synaptically tagged via the ubiquitin-proteasome system (Ageta et al., 2001a,b) and it regulates spinal morphology and synaptic responses (Hennou et al., 2003; Sala et al., 2003). Furthermore, zif268 and vesl-1S knockout mice also exhibit deficient reconsolidation memory processes (see Section IV.E; Bozon et al., 2003; Inoue et al., 2009). These results indicated that neural activitydependent genes have important roles in the memory process. Recently, we examined hippocampal dentate gyrus LTP in urethaneanesthetized rats. Results showed that follistatin or antiactivin A antibody inhibits L-LTP formation without affecting E-LTP (Fig. 9.4A). The decay time course is similar to that of animals injected with the protein synthesis inhibitor anisomycin. Activins facilitate E-LTP duration (Fig. 9.4B) (Ageta et al., 2010), and maintenance of CA1 L-LTP, but not E-LTP, in
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Figure 9.4 Activin is required for L-LTP. (A) Effect of follistatin, antiactivin A, and anisomycin on dentate gyrus LTP persistence in urethane-anesthetized rats. (B) Effect of activin on dentate gyrus LTP persistence in urethane-anesthetized rats. (C) Maintenance of CA1 L-LTP in hippocampal slices from follistatin overexpressing mice and control mice. Two lightning symbols represent strong high-frequency stimulations produced by L-LTP. One lightning symbol represents a weak high-frequency stimulation elicited by E-LTP.
hippocampal slices from follistatin transgenic mice (FBItTA, see below) is significantly reduced compared with control mice (Fig. 9.4C; Ageta et al., 2010). In the marine snail Aplysia, TGF-b induces long-term, but not shortterm, facilitation at synapses between sensory and motor neurons (Zhang et al., 1997). Similarly, treatment with TGF-b2, another isoform of TGF-b, affects synaptic strength and induces CREB phosphorylation in rat cultured hippocampal neurons (Fukushima et al., 2007). Therefore, the TGF-b family of proteins, namely, activins and TGF-b1/2, participate not only in development, but also in neuronal plasticity, of¨ the mature CNS.
E. Activin influences reconsolidation and extinction LTM consists of several distinct processes—acquisition (training), maintenance, and retrieval (recall) phase—through which memory is consolidated. Two recent studies revealed that retrieval of consolidated memory leads to
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two opposing processes: one that weakens old memory and another that strengthens it (Eisenberg et al., 2003; Suzuki et al., 2004). The former process is known as “extinction learning.” The latter process has recently been referred to as “reconsolidation”; memory could be vulnerable following retrieval, so it is reconsolidated in a protein synthesis-dependent manner (Fig. 9.5; Nader, 2003; Nader et al., 2000; Tronson and Taylor, 2007). The study of reconsolidation has extended to numerous learning tasks, such as fear conditioning test (Nader et al., 2000), Morris water maze (Suzuki et al., 2004), and object recognition test (Kelly et al., 2003) in various species such as crabs (Pedreira et al., 2002), chicks (Anokhin et al., 2002), Medaka fish (Eisenberg et al., 2003), rodents (Nader et al., 2000), and humans (Walker et al., 2003). These results suggest that “reconsolidation” is an evolutionarily conserved memory system. Because there is a possibility that enhanced extinction learning or disrupted reconsolidation could be particularly efficacious when treating strong traumatic memory-related disorders, such as posttraumatic stress disorder (PTSD) or phobias, studies have investigated the molecular mechanisms of these processes. In addition, results from our laboratory recently demonstrated that activins in the brain affect both extinction and reconsolidation (Ageta et al., 2010). To examine the role that activin plays in fear memory formation, spatiotemporal-regulated activin (ABItTA) and follistatin (FBItTA) transgenic mice were generated (Ageta et al., 2010). When follistatins are continuously expressed in the brain during training, maintenance, and retrieval phases (Fig. 9.6, Exp. A), FBItTA mice are impaired in LTM but not short-term memory (STM). These results were consistent with activin requirements for L-LTP (see above). Furthermore, a 1-week memory test was performed on FBItTA mice to determine how reconsolidation processes are regulated by activin inhibition (Fig. 9.6, Exp. B). When follistatins were expressed in the brain during maintenance and retrieval phases, there was no significant genotype effect on Test 1 between the FBItTA and control mice (Fig. 9.6, Exp. B). In addition, when the same animals were retested for freezing behavior 24 h later (Test 2 in Exp. B), the FBItTA mice exhibited significantly fewer freezing responses compared with the control mice. When follistatin expression was suppressed during all phases in FBItTA mice, significant genotype effects were not observed in either Test 1 or 2. Therefore, inhibition of activin signals during retrieval resulted in suppressed reconsolidation. Three-week memory testing was also performed on the ABItTA mice, which induces extinction (Fig. 9.7). When activins were upregulated in the brain during maintenance and retrieval phases, there was no significant genotype effect in Test 1 between the ABItTA and control mice (Fig. 9.7, Exp. C). In this experimental paradigm, the freezing level was significantly less in Test 2 compared with Test 1 in control mice. However, there was no significant change between Test 1 and 2 in ABItTA mice.
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Figure 9.5 Memory reconsolidation in 2000, Nader et al. (2000) demonstrated reconsolidation in rats using a cued-fear conditioning test. The illustrations represent the Nader experiment. In brief, rats are placed in chamber A (gray box) and then receive tone and electrical stimulation. After 24 h, rats are placed in chamber B (red columnar box), whose shape is different from that of chamber A, and the rats are exposed to tone (Test 1 in upper row). If the rats associate tone with danger (electrical stimulation), they exhibit freezing behavior. After 24 h, the same rats were retested for freezing behavior (Test 2 in upper line). In the experiment paradigm that triggers reconsolidation process, the rats exhibited freezing behavior again in Test 2 (Test 2 in upper row). In this experimental paradigm, Nader et al. demonstrated that intra-amygdala inhibition of protein synthesis, following retrieval of a previously consolidated memory, resulted in amnesia (no freezing behavior) for the retrieved memory (Test 2 in middle row), but not for consolidated memories that were not retrieved (Test 2 in lower row). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this chapter.)
ABItTA mice also exhibited significantly more freezing than control mice in Test 2. These findings demonstrated that activin upregulation in the brain inhibits the extinction learning.
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Figure 9.6 Contextual fear-conditioning test triggers reconsolidation in FBItTA mice. Utilizing the contextual fear-conditioning test, mice were placed in chamber A (gray box) and were exposed to electrical stimulation. During the retrieval phase, the mice were placed in the same chamber (Test). If the mice associate the chamber with danger, they will exhibit a freezing behavior during the Test phase. In the experiment paradigm that triggers reconsolidation, wild-type mice exhibited freezing behavior again during Test 2 (Test 2 in upper line). In this experimental paradigm, when mice were injected with protein synthesis inhibitor 30 min after Test 1, the mice exhibited reduced freezing behavior in Test 2. In experiment A (Exp. A, middle row), follistatins were continuously expressed during training, maintenance, and retrieval phases in the brain of FBItTA mice. FBItTA mice exhibited reduced freezing behavior during testing in Exp. A. In experiment B (Exp. B, lower row), follistatins were expressed in the brain during maintenance and retrieval phases in FBItTA mice. FBItTA mice exhibited reduced freezing behavior during Test 2, but not during Test 1, of Exp. B. Two lightning symbols represent strong electrical stimulation.
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Figure 9.7 Contextual fear-conditioning test triggers extinction learning in ABItTA mice. In an experiment paradigm that triggers extinction learning, wild-type mice exhibited reduced freezing behavior during Test 2 (Test 2 in upper line). The level of electrical stimulation and length of maintenance phase in Exp. C were different from Exp. B. In experiment C (Exp. C, lower row), activin levels were elevated during maintenance and retrieval phases in the brains of ABItTA mice. ABItTA mice did not exhibit reduced freezing behavior during Test 2 of Exp. C. One lightning symbol represents weak electrical stimulation.
Results from our studies demonstrated that activin inhibition during memory retrieval suppresses previously consolidated fear memories. Therefore, activin signaling could be a promising target for the treatment of disorders that are based on strong traumatic memories, such as PTSD and phobias.
V. Conclusion and Perspectives Activins are involved in various brain functions, including spine formation, anxiety, neurogenesis, L-LTP, LTM, extinction, and reconsolidation (Fig. 9.8). Recent studies have also shown that activins exhibit neurotrophic and neuroprotective effects (Hughes et al., 1999; Tretter et al., 2000; Wu et al., 1999). For these reasons, activin-related compounds have therapeutic potential for candidate drugs to treat CNS disorders. Because antibodies and peptides cannot cross the blood–brain barrier, activins, antiactivin antibodies, and follistatins are not suitable for treating the above-mentioned CNS disorders. Recently, small molecules, such as
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Figure 9.8 Summary of the role of activin in the brain. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this chapter.)
SB-431542 and GW788388, were identified as specific inhibitors of type I activin receptor-like kinase (ALK) receptors, including ActRIB/ALK4, ALK5, and ALK7 (Inman et al., 2002). In addition to small molecule compounds, orally active ALK5 inhibitors were also developed (Gellibert et al., 2006). If these compounds cross the blood–brain barrier, they may also be useful for treating CNS disorders. Because activin-signaling cascades exist in various peripheral tissues, activin-related compounds could result in unpredictable side effects in various tissues. To minimize side effects, it could be essential to combine the development of novel activin-related compounds with brain-specific drug delivery systems. We believe that the development of activin and follistatin transgenic mice could be useful for these drug developments.
ACKNOWLEDGMENTS We acknowledge the collaborative studies with Dr. K. Inokuchi at Toyama University/ MITILS. This work was partly supported by a research grant (H20-018) on psychiatric and neurological diseases from the Ministry of Health, Labour and Welfare and an intramural research grant (20B-13) for neurological and psychiatric disorders of NCNP.
REFERENCES Abdipranoto-Cowley, A., Park, J. S., Croucher, D., Daniel, J., Henshall, S., Galbraith, S., Mervin, K., and Vissel, B. (2009). Activin A is essential for neurogenesis following neurodegeneration. Stem Cells 27, 1330–1346.
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Abraham, W. C., Mason, S. E., Demmer, J., Williams, J. M., Richardson, C. L., Tate, W. P., Lawlor, P. A., and Dragunow, M. (1993). Correlations between immediate early gene induction and the persistence of long-term potentiation. Neuroscience 56, 717–727. Ageta, H., Kato, A., Fukazawa, Y., Inokuchi, K., and Sugiyama, H. (2001a). Effects of proteasome inhibitors on the synaptic localization of Vesl-1S/Homer-1a proteins. Brain Res. Mol. Brain Res. 97, 186–189. Ageta, H., Kato, A., Hatakeyama, S., Nakayama, K., Isojima, Y., and Sugiyama, H. (2001b). Regulation of the level of Vesl-1S/Homer-1a proteins by ubiquitin-proteasome proteolytic systems. J. Biol. Chem. 276, 15893–15897. Ageta, H., Murayama, A., Migishima, R., Kida, S., Tsuchida, K., Yokoyama, M., and Inokuchi, K. (2008). Activin in the brain modulates anxiety-related behavior and adult neurogenesis. PLoS ONE 3, e1869. Ageta, H., Ikegami, S., Miura, M., Masuda, M., Migishima, R., Hino, T., Takashima, N., Murayama, A., Sugino, H., Setou, M., Kida, S., Yokoyama, M., et al. (2010). Activin plays a key role in the maintenance of long-term memory and late-LTP. Learn. Mem. 17, 168–175. Andreasson, K., and Worley, P. F. (1995). Induction of beta-A activin expression by synaptic activity and during neocortical development. Neuroscience 69, 781–796. Anokhin, K. V., Tiunova, A. A., and Rose, S. P. (2002). Reminder effects - reconsolidation or retrieval deficit? Pharmacological dissection with protein synthesis inhibitors following reminder for a passive-avoidance task in young chicks. Eur. J. Neurosci. 15, 1759–1765. Bao, Y. L., Tsuchida, K., Liu, B., Kurisaki, A., Matsuzaki, T., and Sugino, H. (2005). Synergistic activity of activin A and basic fibroblast growth factor on tyrosine hydroxylase expression through Smad3 and ERK1/ERK2 MAPK signaling pathways. J. Endocrinol. 184, 493–504. Bourtchuladze, R., Frenguelli, B., Blendy, J., Cioffi, D., Schutz, G., and Silva, A. J. (1994). Deficient long-term memory in mice with a targeted mutation of the cAMP-responsive element-binding protein. Cell 79, 59–68. Bozon, B., Davis, S., and Laroche, S. (2003). A requirement for the immediate early gene zif268 in reconsolidation of recognition memory after retrieval. Neuron 40, 695–701. Brakeman, P. R., Lanahan, A. A., O’Brien, R., Roche, K., Barnes, C. A., Huganir, R. L., and Worley, P. F. (1997). Homer: A protein that selectively binds metabotropic glutamate receptors. Nature 386, 284–288. Cameron, V. A., Nishimura, E., Mathews, L. S., Lewis, K. A., Sawchenko, P. E., and Vale, W. W. (1994). Hybridization histochemical localization of activin receptor subtypes in rat brain, pituitary, ovary, and testis. Endocrinology 134, 799–808. Castellucci, V. F., Blumenfeld, H., Goelet, P., and Kandel, E. R. (1989). Inhibitor of protein synthesis blocks long-term behavioral sensitization in the isolated gill-withdrawal reflex of Aplysia. J. Neurobiol. 20, 1–9. Cole, A. J., Saffen, D. W., Baraban, J. M., and Worley, P. F. (1989). Rapid increase of an immediate early gene messenger RNA in hippocampal neurons by synaptic NMDA receptor activation. Nature 340, 474–476. de Guise, C., Lacerte, A., Rafiei, S., Reynaud, R., Roy, M., Brue, T., and Lebrun, J. J. (2006). Activin inhibits the human Pit-1 gene promoter through the p38 kinase pathway in a Smad-independent manner. Endocrinology 147, 4351–4362. Di Guglielmo, G. M., Le Roy, C., Goodfellow, A. F., and Wrana, J. L. (2003). Distinct endocytic pathways regulate TGF-beta receptor signalling and turnover. Nat. Cell Biol. 5, 410–421. Dow, A. L., Russell, D. S., and Duman, R. S. (2005). Regulation of activin mRNA and Smad2 phosphorylation by antidepressant treatment in the rat brain: Effects in behavioral models. J. Neurosci. 25, 4908–4916.
Activins Affect Neurogenesis, Anxiety, and Memory
203
Ebisawa, T., Fukuchi, M., Murakami, G., Chiba, T., Tanaka, K., Imamura, T., and Miyazono, K. (2001). Smurf1 interacts with transforming growth factor-beta type I receptor through Smad7 and induces receptor degradation. J. Biol. Chem. 276, 12477–12480. Eisenberg, M., Kobilo, T., Berman, D. E., and Dudai, Y. (2003). Stability of retrieved memory: Inverse correlation with trace dominance. Science 301, 1102–1104. Fischer, M., Kaech, S., Knutti, D., and Matus, A. (1998). Rapid actin-based plasticity in dendritic spines. Neuron 20, 847–854. Fischer, M., Kaech, S., Wagner, U., Brinkhaus, H., and Matus, A. (2000). Glutamate receptors regulate actin-based plasticity in dendritic spines. Nat. Neurosci. 3, 887–894. Frey, U., Krug, M., Reymann, K. G., and Matthies, H. (1988). Anisomycin, an inhibitor of protein synthesis, blocks late phases of LTP phenomena in the hippocampal CA1 region in vitro. Brain Res. 452, 57–65. Fukazawa, Y., Saitoh, Y., Ozawa, F., Ohta, Y., Mizuno, K., and Inokuchi, K. (2003). Hippocampal LTP is accompanied by enhanced F-actin content within the dendritic spine that is essential for late LTP maintenance in vivo. Neuron 38, 447–460. Fukushima, T., Liu, R. Y., and Byrne, J. H. (2007). Transforming growth factor-beta2 modulates synaptic efficacy and plasticity and induces phosphorylation of CREB in hippocampal neurons. Hippocampus 17, 5–9. Funaba, M., Murata, T., Fujimura, H., Murata, E., Abe, M., and Torii, K. (1997). Immunolocalization of type I or type II activin receptors in the rat brain. J. Neuroendocrinol. 9, 105–111. Gage, F. H. (2002). Neurogenesis in the adult brain. J. Neurosci. 22, 612–613. Gamer, L. W., Wolfman, N. M., Celeste, A. J., Hattersley, G., Hewick, R., and Rosen, V. (1999). A novel BMP expressed in developing mouse limb, spinal cord, and tail bud is a potent mesoderm inducer in Xenopus embryos. Dev. Biol. 208, 222–232. Gellibert, F., de Gouville, A. C., Woolven, J., Mathews, N., Nguyen, V. L., BerthoRuault, C., Patikis, A., Grygielko, E. T., Laping, N. J., and Huet, S. (2006). Discovery of 4-{4-[3-(pyridin-2-yl)-1H-pyrazol-4-yl]pyridin-2-yl}-N-(tetrahydro-2H- pyran-4yl)benzamide (GW788388): A potent, selective, and orally active transforming growth factor-beta type I receptor inhibitor. J. Med. Chem. 49, 2210–2221. Hata, Y., and Takai, Y. (1999). Roles of postsynaptic density-95/synapse-associated protein 90 and its interacting proteins in the organization of synapses. Cell. Mol. Life Sci. 56, 461–472. Hennou, S., Kato, A., Schneider, E. M., Lundstrom, K., Gahwiler, B. H., Inokuchi, K., Gerber, U., and Ehrengruber, M. U. (2003). Homer-1a/Vesl-1S enhances hippocampal synaptic transmission. Eur. J. Neurosci. 18, 811–819. Honkura, N., Matsuzaki, M., Noguchi, J., Ellis-Davies, G. C., and Kasai, H. (2008). The subspine organization of actin fibers regulates the structure and plasticity of dendritic spines. Neuron 57, 719–729. Hughes, P. E., Alexi, T., Williams, C. E., Clark, R. G., and Gluckman, P. D. (1999). Administration of recombinant human Activin-A has powerful neurotrophic effects on select striatal phenotypes in the quinolinic acid lesion model of Huntington’s disease. Neuroscience 92, 197–209. Inman, G. J., Nicolas, F. J., Callahan, J. F., Harling, J. D., Gaster, L. M., Reith, A. D., Laping, N. J., and Hill, C. S. (2002). SB-431542 is a potent and specific inhibitor of transforming growth factor-beta superfamily type I activin receptor-like kinase (ALK) receptors ALK4, ALK5, and ALK7. Mol. Pharmacol. 62, 65–74. Inokuchi, K., Kato, A., Hiraia, K., Hishinuma, F., Inoue, M., and Ozawa, F. (1996). Increase in activin beta A mRNA in rat hippocampus during long-term potentiation. FEBS Lett. 382, 48–52. Inoue, N., Nakao, H., Migishima, R., Hino, T., Matsui, M., Hayashi, F., Nakao, K., Manabe, T., Aiba, A., and Inokuchi, K. (2009). Requirement of the immediate early gene vesl-1S/homer-1a for fear memory formation. Mol. Brain 2, 7.
204
Hiroshi Ageta and Kunihiro Tsuchida
Jones, M. W., Errington, M. L., French, P. J., Fine, A., Bliss, T. V., Garel, S., Charnay, P., Bozon, B., Laroche, S., and Davis, S. (2001). A requirement for the immediate early gene Zif268 in the expression of late LTP and long-term memories. Nat. Neurosci. 4, 289–296. Kato, A., Ozawa, F., Saitoh, Y., Hirai, K., and Inokuchi, K. (1997). vesl, a gene encoding VASP/Ena family related protein, is upregulated during seizure, long-term potentiation and synaptogenesis. FEBS Lett. 412, 183–189. Kato, A., Ozawa, F., Saitoh, Y., Fukazawa, Y., Sugiyama, H., and Inokuchi, K. (1998). Novel members of the Vesl/Homer family of PDZ proteins that bind metabotropic glutamate receptors. J. Biol. Chem. 273, 23969–23975. Kelly, A., Laroche, S., and Davis, S. (2003). Activation of mitogen-activated protein kinase/ extracellular signal-regulated kinase in hippocampal circuitry is required for consolidation and reconsolidation of recognition memory. J. Neurosci. 23, 5354–5360. Kessler, R. C., McGonagle, K. A., Zhao, S., Nelson, C. B., Hughes, M., Eshleman, S., Wittchen, H. U., and Kendler, K. S. (1994). Lifetime and 12-month prevalence of DSMIII-R psychiatric disorders in the United States. Results from the National Comorbidity Survey. Arch. Gen. Psychiatry 51, 8–19. Kitamura, T., Saitoh, Y., Takashima, N., Murayama, A., Niibori, Y., Ageta, H., Sekiguchi, M., Sugiyama, H., and Inokuchi, K. (2009). Adult neurogenesis modulates the hippocampus-dependent period of associative fear memory. Cell 139, 814–827. Kokaia, Z., and Lindvall, O. (2003). Neurogenesis after ischaemic brain insults. Curr. Opin. Neurobiol. 13, 127–132. Kuhn, H. G., Dickinson-Anson, H., and Gage, F. H. (1996). Neurogenesis in the dentate gyrus of the adult rat: Age-related decrease of neuronal progenitor proliferation. J. Neurosci. 16, 2027–2033. Kurisaki, A., Inoue, I., Kurisaki, K., Yamakawa, N., Tsuchida, K., and Sugino, H. (2008). Activin induces long-lasting N-methyl-D-aspartate receptor activation via scaffolding PDZ protein activin receptor interacting protein 1. Neuroscience 151, 1225–1235. Ling, N., Ying, S. Y., Ueno, N., Shimasaki, S., Esch, F., Hotta, M., and Guillemin, R. (1986). Pituitary FSH is released by a heterodimer of the beta-subunits from the two forms of inhibin. Nature 321, 779–782. Lyford, G. L., Yamagata, K., Kaufmann, W. E., Barnes, C. A., Sanders, L. K., Copeland, N. G., Gilbert, D. J., Jenkins, N. A., Lanahan, A. A., and Worley, P. F. (1995). Arc, a growth factor and activity-regulated gene, encodes a novel cytoskeletonassociated protein that is enriched in neuronal dendrites. Neuron 14, 433–445. Malberg, J. E., Eisch, A. J., Nestler, E. J., and Duman, R. S. (2000). Chronic antidepressant treatment increases neurogenesis in adult rat hippocampus. J. Neurosci. 20, 9104–9110. Mather, J. P., Moore, A., and Li, R. H. (1997). Activins, inhibins, and follistatins: Further thoughts on a growing family of regulators. Proc. Soc. Exp. Biol. Med. 215, 209–222. Matus, A. (2005). Growth of dendritic spines: A continuing story. Curr. Opin. Neurobiol. 15, 67–72. Matzuk, M. M., Kumar, T. R., and Bradley, A. (1995a). Different phenotypes for mice deficient in either activins or activin receptor type II. Nature 374, 356–360. Matzuk, M. M., Kumar, T. R., Vassalli, A., Bickenbach, J. R., Roop, D. R., Jaenisch, R., and Bradley, A. (1995b). Functional analysis of activins during mammalian development. Nature 374, 354–356. McPherron, A. C., Lawler, A. M., and Lee, S. J. (1997). Regulation of skeletal muscle mass in mice by a new TGF-beta superfamily member. Nature 387, 83–90. Muir, K. W. (2006). Glutamate-based therapeutic approaches: Clinical trials with NMDA antagonists. Curr. Opin. Pharmacol. 6, 53–60. Mukherjee, S., Sackeim, H. A., and Schnur, D. B. (1994). Electroconvulsive therapy of acute manic episodes: A review of 50 years’ experience. Am. J. Psychiatry 151, 169–176. Nader, K. (2003). Memory traces unbound. Trends Neurosci. 26, 65–72.
Activins Affect Neurogenesis, Anxiety, and Memory
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Nader, K., Schafe, G. E., and Le Doux, J. E. (2000). Fear memories require protein synthesis in the amygdala for reconsolidation after retrieval. Nature 406, 722–726. Nakamura, T., Asashima, M., Eto, Y., Takio, K., Uchiyama, H., Moriya, N., Ariizumi, T., Yashiro, T., Sugino, K., Titani, K., et al. (1992). Isolation and characterization of native activin B. J. Biol. Chem. 267, 16385–16389. Nakashima, M., Toyono, T., Akamine, A., and Joyner, A. (1999). Expression of growth/ differentiation factor 11, a new member of the BMP/TGFbeta superfamily during mouse embryogenesis. Mech. Dev. 80, 185–189. Nguyen, P. V., Abel, T., and Kandel, E. R. (1994). Requirement of a critical period of transcription for induction of a late phase of LTP. Science 265, 1104–1107. Pangas, S. A., and Woodruff, T. K. (2000). Activin signal transduction pathways. Trends Endocrinol. Metab. 11, 309–314. Parsons, C. G., Danysz, W., and Quack, G. (1999). Memantine is a clinically well tolerated N-methyl-D-aspartate (NMDA) receptor antagonist—A review of preclinical data. Neuropharmacology 38, 735–767. Pedreira, M. E., Perez-Cuesta, L. M., and Maldonado, H. (2002). Reactivation and reconsolidation of long-term memory in the crab Chasmagnathus: Protein synthesis requirement and mediation by NMDA-type glutamatergic receptors. J. Neurosci. 22, 8305–8311. Plath, N., Ohana, O., Dammermann, B., Errington, M. L., Schmitz, D., Gross, C., Mao, X., Engelsberg, A., Mahlke, C., Welzl, H., Kobalz, U., Stawrakakis, A., et al. (2006). Arc/ Arg3.1 is essential for the consolidation of synaptic plasticity and memories. Neuron 52, 437–444. Sala, C., Futai, K., Yamamoto, K., Worley, P. F., Hayashi, Y., and Sheng, M. (2003). Inhibition of dendritic spine morphogenesis and synaptic transmission by activity-inducible protein Homer1a. J. Neurosci. 23, 6327–6337. Santarelli, L., Saxe, M., Gross, C., Surget, A., Battaglia, F., Dulawa, S., Weisstaub, N., Lee, J., Duman, R., Arancio, O., Belzung, C., and Hen, R. (2003). Requirement of hippocampal neurogenesis for the behavioral effects of antidepressants. Science 301, 805–809. Sawa, A., and Snyder, S. H. (2002). Schizophrenia: Diverse approaches to a complex disease. Science 296, 692–695. Sekiguchi, M., Hayashi, F., Tsuchida, K., and Inokuchi, K. (2009). Neuron type-selective effects of activin on development of the hippocampus. Neurosci. Lett. 452, 232–237. Sharma, M., Kambadur, R., Matthews, K. G., Somers, W. G., Devlin, G. P., Conaglen, J. V., Fowke, P. J., and Bass, J. J. (1999). Myostatin, a transforming growth factor-beta superfamily member, is expressed in heart muscle and is upregulated in cardiomyocytes after infarct. J. Cell. Physiol. 180, 1–9. Sheng, M., and Sala, C. (2001). PDZ domains and the organization of supramolecular complexes. Annu. Rev. Neurosci. 24, 1–29. Shoji, H., Tsuchida, K., Kishi, H., Yamakawa, N., Matsuzaki, T., Liu, Z., Nakamura, T., and Sugino, H. (2000). Identification and characterization of a PDZ protein that interacts with activin type II receptors. J. Biol. Chem. 275, 5485–5492. Shoji-Kasai, Y., Ageta, H., Hasegawa, Y., Tsuchida, K., Sugino, H., and Inokuchi, K. (2007). Activin increases the number of synaptic contacts and the length of dendritic spine necks by modulating spinal actin dynamics. J. Cell Sci. 120, 3830–3837. Shors, T. J., Miesegaes, G., Beylin, A., Zhao, M., Rydel, T., and Gould, E. (2001). Neurogenesis in the adult is involved in the formation of trace memories. Nature 410, 372–376. Skolnick, P. (1999). Antidepressants for the new millennium. Eur. J. Pharmacol. 375, 31–40. Squire, L. R., and Barondes, S. H. (1973). Memory impairment during prolonged training in mice given inhibitors of cerebral protein synthesis. Brain Res. 56, 215–225. Suzuki, A., Josselyn, S. A., Frankland, P. W., Masushige, S., Silva, A. J., and Kida, S. (2004). Memory reconsolidation and extinction have distinct temporal and biochemical signatures. J. Neurosci. 24, 4787–4795.
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Hiroshi Ageta and Kunihiro Tsuchida
Talmage, D. A. (2008). Mechanisms of neuregulin action. Novartis Found. Symp. 289, 74–84, discussion 84–93. ten Dijke, P., and Hill, C. S. (2004). New insights into TGF-beta-Smad signalling. Trends Biochem. Sci. 29, 265–273. ten Dijke, P., Miyazono, K., and Heldin, C. H. (2000). Signaling inputs converge on nuclear effectors in TGF-beta signaling. Trends Biochem. Sci. 25, 64–70. Tretter, Y. P., Hertel, M., Munz, B., ten Bruggencate, G., Werner, S., and Alzheimer, C. (2000). Induction of activin A is essential for the neuroprotective action of basic fibroblast growth factor in vivo. Nat. Med. 6, 812–815. Tronson, N. C., and Taylor, J. R. (2007). Molecular mechanisms of memory reconsolidation. Nat. Rev. Neurosci. 8, 262–275. Tsuchida, K. (2004). Activins, myostatin and related TGF-beta family members as novel therapeutic targets for endocrine, metabolic and immune disorders. Curr. Drug Targets Immune Endocr. Metabol. Disord. 4, 157–166. Tsuchida, K., Nakatani, M., Hitachi, K., Uezumi, A., Sunada, Y., Ageta, H., and Inokuchi, K. (2009). Activin signaling as an emerging target for therapeutic interventions. Cell Commun. Signal 7, 15. Tu, J. C., Xiao, B., Naisbitt, S., Yuan, J. P., Petralia, R. S., Brakeman, P., Doan, A., Aakalu, V. K., Lanahan, A. A., Sheng, M., and Worley, P. F. (1999). Coupling of mGluR/Homer and PSD-95 complexes by the Shank family of postsynaptic density proteins. Neuron 23, 583–592. Vale, W., Rivier, J., Vaughan, J., McClintock, R., Corrigan, A., Woo, W., Karr, D., and Spiess, J. (1986). Purification and characterization of an FSH releasing protein from porcine ovarian follicular fluid. Nature 321, 776–779. Walker, M. P., Brakefield, T., Hobson, J. A., and Stickgold, R. (2003). Dissociable stages of human memory consolidation and reconsolidation. Nature 425, 616–620. Wang, R., Dineley, K. T., Sweatt, J. D., and Zheng, H. (2004). Presenilin 1 familial Alzheimer’s disease mutation leads to defective associative learning and impaired adult neurogenesis. Neuroscience 126, 305–312. Werner, S., and Alzheimer, C. (2006). Roles of activin in tissue repair, fibrosis, and inflammatory disease. Cytokine Growth Factor Rev. 17, 157–171. Wong, M. L., and Licinio, J. (2004). From monoamines to genomic targets: A paradigm shift for drug discovery in depression. Nat. Rev. Drug Discov. 3, 136–151. Wu, D. D., Lai, M., Hughes, P. E., Sirimanne, E., Gluckman, P. D., and Williams, C. E. (1999). Expression of the activin axis and neuronal rescue effects of recombinant activin A following hypoxic-ischemic brain injury in the infant rat. Brain Res. 835, 369–378. Wu, H. H., Ivkovic, S., Murray, R. C., Jaramillo, S., Lyons, K. M., Johnson, J. E., and Calof, A. L. (2003). Autoregulation of neurogenesis by GDF11. Neuron 37, 197–207. Yamaguchi, T., Kurisaki, A., Yamakawa, N., Minakuchi, K., and Sugino, H. (2006). FKBP12 functions as an adaptor of the Smad7-Smurf1 complex on activin type I receptor. J. Mol. Endocrinol. 36, 569–579. Yao, I., Takagi, H., Ageta, H., Kahyo, T., Sato, S., Hatanaka, K., Fukuda, Y., Chiba, T., Morone, N., Yuasa, S., Inokuchi, K., Ohtsuka, T., et al. (2007). SCRAPPER-dependent ubiquitination of active zone protein RIM1 regulates synaptic vesicle release. Cell 130, 943–957. Ying, S. Y., Zhang, Z., Furst, B., Batres, Y., Huang, G., and Li, G. (1997). Activins and activin receptors in cell growth. Proc. Soc. Exp. Biol. Med. 214, 114–122. Zhang, F., Endo, S., Cleary, L. J., Eskin, A., and Byrne, J. H. (1997). Role of transforming growth factor-beta in long-term synaptic facilitation in Aplysia. Science 275, 1318–1320. Zheng, F., Adelsberger, H., Muller, M. R., Fritschy, J. M., Werner, S., and Alzheimer, C. (2009). Activin tunes GABAergic neurotransmission and modulates anxiety-like behavior. Mol. Psychiatry 14, 332–346.
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The Role of Activin/Nodal and Wnt Signaling in Endoderm Formation Catherine Payne,* Jason King,† and David Hay* Contents 208 209 210 212 214 214 214
I. Introduction II. TGFb Signaling III. WNT/b-Catenin Signaling IV. Applied Biology V. Conclusions Acknowledgments References
Abstract Human embryonic stem cells (hESCs) are located within the inner cell mass of the preimplantation blastocysts. hESCs exhibit two important properties, the ability to generate exact copies of themselves, termed self-renewal, and pluripotency, the ability of stem cells to differentiate into every cell type of the embryo. This means that in theory it may be possible to generate an inexhaustible supply of primary human somatic cells in vitro which are suitable for application in regenerative medicine. Maintaining stem cell self-renewal and eliciting differentiation are dependent on the coordination of a number of signaling pathways which include members of the transforming growth factor beta (TGFb) and Wnt families. The work in our laboratory has focused on the efficient generation of hepatocyte-like cells (HLCs) from hESCs and induced pluripotent stem cells (iPSCs). In order to mimic signaling during primitive streak and endoderm development, we have utilized TGFb and Wnt signaling pathways in vitro. This has resulted in the generation of homogeneous populations of HLCs exhibiting liver specific function. This chapter will focus on TGFb and Wnt signaling pathways and their role in primitive streak, endoderm, and HLC development. ß 2011 Elsevier Inc.
* MRC Centre for Regenerative Medicine, The University of Edinburgh, Edinburgh, United Kingdom Roslin Cellab, Roslin Biocentre, Roslin, Midlothian, Scotland, United Kingdom
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Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00010-X
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I. Introduction Human development is a complex process involving the integration of multiple signaling pathways regulating gene transcription. These pathways are essential for patterning the embryo and eliciting differentiation to all the three germ layers: endoderm, mesoderm, and ectoderm (Tam and Loebel, 2007). In particular, we will focus on the generation of definitive endoderm (DE) as a precursor to HLCs. DE differentiation is preceded by the ingression of the epiblast (primitive ectoderm) to form the primitive steak (PS) from which both endoderm and mesoderm arise (Ginsburg et al., 1990; Tam and Loebel, 2007; Tam et al., 2004). The two major factors involved in PS formation, Activin/Nodal and Wnt, also play important roles in endoderm and mesoderm specification (Ding et al., 1998; Gadue et al., 2006; Liu et al., 1999; Yamamoto et al., 2001). Following commitment, DE lines the ventral region of the developing embryo forming the foregut, midgut, and hindgut (Kaestner, 2005; Zhao and Duncan, 2005). The foregut endoderm develops into the lung, thyroid, liver, and the ventral rudiment of the pancreas and is patterned by factors secreted from the adjacent mesenchymal structures (Dessimoz and Grapin-Botton, 2006; Lemaigre and Zaret, 2004; Zaret, 2001) in a concentration-dependent manner (Serls et al., 2005). The region of the foregut which becomes the liver bud is adjacent to the cardiac mesoderm (CM) and the septum transversum (ST). Both structures secrete factors which pattern the foregut endoderm (the CM produces FGFs while the ST produces BMPs) (Douarin, 1975; Rossi et al., 2001). In vitro, hESC-derived DE can be differentiated to hepatic endoderm using low levels of FGF2 while intermediate and high concentrations induces a pancreatic cell fate and pulmonary fate, respectively (Ameri et al., 2010). Following specification, the emerging liver bud invades the ST and the resident liver stem cell, the hepatoblast, differentiates into two of the major cell types found in the liver, hepatocytes and cholangiocytes (for a review, see Dancygier, 2010). We have studied human development in vitro and in vivo and applied this knowledge to pluripotent stem cells. In doing so, we have generated a system for deriving large numbers of high fidelity HLCs in vitro. These HLCs exhibit many of the attributes of primary human hepatocytes and will play important roles in developing cell-based assays which model human drug toxicity and disease. Additionally an inexhaustible supply of HLCs has an important role to play in developing cell-based therapies for human liver disease (for a review, see Dalgetty et al., 2009).
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II. TGFb Signaling TGFb signaling plays an important role in embryonic development and involves the interplay of a large number of ligands, regulators, and receptors (Heldin et al., 2009; Kitisin et al., 2007; Fig. 10.1). The TGFb superfamily are soluble ligands, including nodal, Activin A, BMPs, and
Cerberus
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Figure 10.1 Smad-mediated pathway of Activin/Nodal and BMP signaling. Activin binding (1) to the Activin receptor (ActR) type II dimer induces coassociation of a type I Activin receptor dimer (in the case of Activin, this is ALK4) which leads to the phosphorylation and activation of ALK4 by the type II receptor’s kinase (3). The Smad 2/3 complex is recruited to ALK4 (4) via interaction with SARA. Phosphorylation of Smad 2/3 (5) results in a conformational change that enables it to bind Smad 4 (7). This complex enters the nucleus (8) and associates with transcriptional cofactors (9) and P300/CBP to drive expression (10). Prior to Nodal binding, a member of the EGFCFC coreceptor family, such as Cripto, must first bind to ALK4 to stabilize the tetrameric ALK4-ActRII receptor complex. Subsequent signaling is via the same Smad 2/3 pathway. BMP signaling also requires assembly of a tetrameric receptor complex; however, these can be more varied involving an additional type II receptor (BMPRII) and three different type I receptors (ALK2, ALK3, and ALK6). Instead of using Smad 2/3þ4, BMP signaling can use Smads 1/5/8þ4. Nodal signaling can be inhibited extracellularly by Lefty and Cerberus. Inhibitory Smads 6 and 7 are strongly induced by Activin, TGF-b, and BMP and act as intracellular antagonists of Smad-mediated signaling.
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TGFb (Kitisin et al., 2007), which bind to type II and type I receptors. Nodal/Activin A bind to the extracellular portion of the Activin receptor type 2 (ActRIIA or ActRIIB) dimer, causing a conformational change and phosphorylation of the glycine serine rich domain (GS) on the Activin receptor type 1B (ALK4). BMPs also bind to several type II receptors (ActRIIA, ActRIIB, and BMPR2) activating the type I receptor (ALK2, ALK3, or ALK6; Heldin et al., 1997; Kitisin et al., 2007; Moustakas, 2002; Fig. 10.1). Following activation, the type I receptor recruits the receptorregulated Smads to the receptor complex through the SMAD anchor for receptor activation (SARA). The activated type I receptors then phosphorylate the intracellular messenger Smads (Heldin et al., 1997). There are at least eight Smad proteins which can be split into three classes: (i) receptoractivated Smads (R-Smads) include Smad 1, Smad 2, Smad 3, Smad 5, and Smad 8/9; (ii) the comediator or common Smad (co-Smad), Smad 4; and (iii) inhibitory Smads (I-Smads) include Smad 6 and Smad 20–22. Following Activin/Nodal signaling, Smads 2 and 3 dissociate from SARA and bind Smad 4. The Smad 2/3/4 complex then translocates to the nucleus and activates gene expression (Fig. 10.1). BMP signaling induces the activation of Smad 1, 5, or 8 which bind to Smad 4 and translocate to the nucleus and regulate gene transcription (Feng and Derynck, 2005; Heldin et al., 1997; Itoh and ten Dijke, 2007; Kitisin et al., 2007; Massague et al., 2005; Moustakas, 2002; Fig. 10.1). Activin/Nodal signaling plays a central role in patterning of vertebrate embryos and in vitro Activin A binds the same receptor as Nodal and likely mimics Nodal signaling (Gadue et al., 2006). The role of Nodal signaling in the mouse during development has been studied using insertional mutagenesis and receptor complex nulls. Both types of mutants failed to form an elongated primitive streak, displaying excessive ectoderm commitment and deficiency in endoderm and mesoderm. This was in part rescued by hypomorphic expression of Nodal revealing a degree of DE differentiation (Lowe et al., 2001). The requirement of high concentrations of Activin/ Nodal signaling in the formation of the primitive streak and subsequently DE was further supported by studies in hESCs (D’Amour et al., 2005) demonstrating a pivotal role of TGFb signaling in human endoderm development.
III. WNT/b-Catenin Signaling Wnt signaling also plays an important role in embryonic development. The Wnt signaling pathway is a complex network involving a large number of ligands, regulators, and receptors (Logan and Nusse, 2004; Moon et al., 1997; Fig. 10.2). There are at least 19 Wnt genes and 10 frizzled receptors
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Figure 10.2 Wnt/b-Catenin signaling pathway. In the absence of Wnt, LRP is bound by Dickkopf (DKK) and prevented from interacting with Frizzled. b-Catenin is bound by CK1a, Axin, GSK3b, and APC and phosphorylated to establish the “Destruction ¨ b-Catenin to the proteasome for degradation. When Wnt Complex” which targets binds Frizzled, LRP and Disheveled (DVL) associate with Frizzled and promote the dissociation of the destruction complex, to release APC and enable the remaining proteins to form a complex with LRP/Frizzled/DVL. b-Catenin is then free to translocate to the nucleus where it displaces the GRO/HDAC/CTBP repression complex from the TCF (T cell factor)-promoting expression.
which are part of the G protein-coupled receptor family which can be broadly divided into canonical and noncanonical signaling pathways. The canonical Wnt pathway involves the binding of Wnt to its cell-surface receptor, Frizzled, and its coreceptor low density lipoprotein receptor protein (LRP5/6). This causes the activation of the Dishevelled (DSH) family proteins, which in turn recruits axin to the plasma membrane and inhibits the assembly of the destruction complex (Axin, GSK3b APC, and CKI) which promotes b-Catenin stability and subsequent nuclear translocation (Logan and Nusse, 2004; Moon et al., 1997). In the nucleus, b-Catenin interacts with the T cell factor/lymphocyte enhancing binding factor (TCF/LEF) family of transcription factors displacing the transcriptional repressor Groucho and HDAC (Fig. 10.2; Arce et al., 2006; Clevers, 2006; Moon, 2005; van Noort and Clevers, 2002; Wu and Nusse, 2002). In the absence of Wnt signaling, b-Catenin is tagged for degradation by the
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“destruction complex” and does not lead to transcriptional activation by b-Catenin (Clevers, 2006; Kimelman and Xu, 2006). There are other Wnt pathways which do not involve the frizzled receptor/b-Catenin pathway and are referred to as noncanonical. This signaling pathway either elicits an increase in intracellular calcium through interactions with protein kinase C or activates small Rho GTPases, such as c-Jun N terminus kinase ( Jnk) (Kuhl et al., 2000; Nateri et al., 2005; Topol et al., 2003; Westfall et al., 2003). For the simplicity, we will concentrate on canonical signaling and its role in PS formation. Canonical Wnt signaling has been shown to play important role in PS and endoderm development through knockout studies. The effects of knocking out the intracellular messenger, b-Catenin, have been studied in mice. While b-Catenin null embryos formed blastocysts that implanted and developed to egg cylinder stage embryos, they did not form the primitive streak, mesoderm, or endoderm (Haegel et al., 1995). Further work in the field demonstrated that Wnt3 was essential for PS formation demonstrating the importance of Wnt3 in primary axis formation in the mammals (Liu et al., 1999). More recently, canonical Wnt signaling was shown to be essential in the proper development of the liver bud in zebra fish (Ober et al., 2006). In support of this, canonical Wnt signaling has been shown important in human liver development and hESC differentiation to both PS and HLCs (Hay et al., 2008).
IV. Applied Biology The coordinated expression and function of both Activin/Nodal and Wnt are essential for proper PS and derivative cell-type formation in both mouse and human development (Gadue et al., 2006; Hay et al., 2008). When coupled to pluripotent stem cells, it provides the biologist with an enormous potential to generate efficient levels of endoderm or mesoderm derivatives for in vitro and in vivo experimentation. In this section, we will discuss the importance of this technology with a focus on the generation of HLCs from pluripotent stem cell populations via the PS. The real impact of deriving hepatocyte-like cells (HLCs) from pluripotent stem cell is the provision of a resource from the desired genetic background. The current gold standard are primary human hepatocytes which are human in origin and possess broad range function upon isolation. Unfortunately, high-level liver function is lost within 24–48 h of isolation. Other limitations include limited supply and expense. Alternative approaches have employed primary rat, porcine, and murine hepatocytes which are useful, but often not indicative of human liver function and also possess limited clinical application. Transformed human hepatocytes have
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also been used to model human liver function and while the lines can be propagated in vitro, they exhibit poor differentiated function which limits their use (Sharma et al., 2010). Therefore the provision of high fidelity HLCs from a renewable and scalable source, such as hESC or iPSC, would benefit a number of downstream regenerative medical applications playing a role in society’s health and wealth. The accurate prediction of human drug toxicity is one example. The costs of successful drug development are heavily influenced by compound attrition rate. For every new drug that reaches the market, 5000–10,000 compounds have been tested in preclinical trials. Therefore, any strategies which streamline and standardize the process of predictive toxicology testing would impact drug attrition levels and therefore cost. Stem cell-derived liver technology could also be applied to the creation of models which allow us to model “disease in a dish.” The recent advances in cellular reprogramming of human somatic cells have allowed us to generate iPSCs from human somatic cells (for a review, see Dalgetty et al., 2009). This in essence has given us the ability to create human libraries of stem cells which display the genetic background of interest. We have shown that it is possible to translate the know-how developed in hESCs and apply that to iPSCs to efficiently deriving hepatocytes (Sullivan et al., 2010) which hold great potential in developing a better understanding of human liver toxicity and disease. Presently, end stage liver disease is on the increase in the Western world. Although highly successful, orthotopic liver transplantation is not the answer to this problem due to limited availability of organs. This is compounded by the dire prediction that these incidences to liver disease will rise sharply in the next 10 years. As such, there is an urgent need to develop alternative treatment strategies. One example of this is the bioartificial liver (BAL) device, which is viewed as a pragmatic approach to this problem (for a review, see Carpentier et al., 2009). While a number of groups have developed promising technologies, the ability to translate those to a cost-effective good manufacturing practice (GMP) BAL is proving difficult. This is mainly due to the numerous biological issues associated with primary hepatocytes and cell lines and may benefit from large-scale production of hESC- or iPSC-derived HLCs. Another approach to treating liver disease, and probably the most challenging, is the provision of functional mass by cell transplantation. Encouraging experiments have shown previously that primary adult hepatocytes successfully transplant into diseased organs resulting in a modest benefit (Bilir et al., 2000; Strom et al., 1997). However, the issues of primary human hepatocyte scarcity and heterogeneity mean that this approach is not feasible in the clinic. While hESC HLCs offer great potential here, there are numerous cell culture challenges that face this strategy, such as their defined, cost-effective, and GMP scalable production (Hannoun et al., 2010). Probably, the greatest challenge that faces cell therapy in the liver and other tissues is the safe transplantation of the derivative cells without tumor formation in vivo (Basma et al., 2009).
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V. Conclusions Applying the knowledge acquired from a number of developmental studies has proved vital in developing high fidelity somatic cell models from pluripotent stem cells. In particular, we have studied Activin/Nodal and Wnt pathways which play an important role in primitive streak, DE, and HLC formation from both hESC and iPSCs. As such, pluripotent stem cellderived HLCs are likely to revolutionize the way in which we model human liver development and disease, measure human drug toxicity, and provide novel cell therapies for treating human disease.
ACKNOWLEDGMENTS Drs. Payne and King were funded by a grant from the UK Stem Cell Foundation and Scottish Enterprise. Dr. Hay was funded by a RCUK Fellowship.
REFERENCES Ameri, J., Stahlberg, A., Pedersen, J., Johansson, J. K., Johannesson, M. M., Artner, I., et al. (2010). FGF2 specifies hESC-derived definitive endoderm into foregut/midgut cell lineages in a concentration-dependent manner. Stem Cells 28(1), 45–56. Arce, L., Yokoyama, N. N., and Waterman, M. L. (2006). Diversity of LEF/TCF action in development and disease. Oncogene 25(57), 7492–7504. Basma, H., Soto-Gutie´rrez, A., Yannam, G. R., Liu, L., Ito, R., Yamamoto, T., et al. (2009). Differentiation and transplantation of human embryonic stem cell-derived hepatocytes. Gastroenterology 136(3), 990–999. Bilir, B. M., Guinette, D., Karrer, F., Kumpe, D. A., Krysl, J., Stephens, J., et al. (2000). Hepatocyte transplantation in acute liver failure. Liver Transpl. 6(1), 32–40. Carpentier, B., Gautier, A., and Legallais, C. (2009). Artificial and bio-artificial liver devices: Present and future. Gut 58(12), 1690–1702. Clevers, H. (2006). Wnt/beta-catenin signaling in development and disease. Cell 127(3), 469–480. Dalgetty, D. M., Medine, C. N., Iredale, J. P., and Hay, D. C. (2009). Progress and future challenges in stem cell-derived liver technologies. Am. J. Physiol. Gastrointest. Liver Physiol. 297(2), G241–G248. D’Amour, K. A., Agulnick, A. D., Eliazer, S., Kelly, O. G., Kroon, E., and Baetge, E. E. (2005). Efficient differentiation of human embryonic stem cells to definitive endoderm. Nat. Biotechnol. 23(12), 1534–1541. Dancygier, H. (2010). Clinical hepatology: Principles and practice of hepatobiliary diseases. Springer Verlag. Dessimoz, J., and Grapin-Botton, A. (2006). Pancreas development and cancer: Wnt/betacatenin at issue. Cell Cycle 5(1), 7–10. Ding, J., Yang, L., Yan, Y. T., Chen, A., Desai, N., Wynshaw-Boris, A., et al. (1998). Cripto is required for correct orientation of the anterior-posterior axis in the mouse embryo. Nature 395(6703), 702–707.
The Role of Activin/Nodal and Wnt Signaling in Endoderm Formation
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Douarin, N. M. (1975). An experimental analysis of liver development. Med. Biol. 53(6), 427–455. Feng, X. H., and Derynck, R. (2005). Specificity and versatility in tgf-beta signaling through Smads. Annu. Rev. Cell Dev. Biol. 21, 659–693. Gadue, P., Huber, T. L., Paddison, P. J., and Keller, G. M. (2006). Wnt and TGF-beta signaling are required for the induction of an in vitro model of primitive streak formation using embryonic stem cells. Proc. Natl. Acad. Sci. USA 103(45), 16806–16811. Ginsburg, M., Snow, M. H., and McLaren, A. (1990). Primordial germ cells in the mouse embryo during gastrulation. Development 110(2), 521–528. Haegel, H., Larue, L., Ohsugi, M., Fedorov, L., Herrenknecht, K., and Kemler, R. (1995). Lack of beta-catenin affects mouse development at gastrulation. Development 121(11), 3529–3537. Hannoun, Z., Fletcher, J., Greenhough, S., Medine, C. N., Samuel, K., Sharma, R., Pryde, A., et al. (2010). The comparison between conditioned media and serum free media in human embryonic stem cell culture and differentiation. Cell. Reprogram. 12(2), 133–140. Hay, D. C., Fletcher, J., Payne, C., Terrace, J. D., Gallagher, R. C., Snoeys, J., et al. (2008). Highly efficient differentiation of hESCs to functional hepatic endoderm requires ActivinA and Wnt3a signaling. Proc. Natl. Acad. Sci. USA 105(34), 12301–12306. Heldin, C. H., Miyazono, K., and Ten, D. P. (1997). TGF-beta signalling from cell membrane to nucleus through SMAD proteins. Nature 390(6659), 465–471. Heldin, C. H., Landstrom, M., and Moustakas, A. (2009). Mechanism of TGF-beta signaling to growth arrest, apoptosis, and epithelial-mesenchymal transition. Curr. Opin. Cell Biol. 21(2), 166–176. Itoh, S., and ten Dijke, P. (2007). Negative regulation of TGF-beta receptor/Smad signal transduction. Curr. Opin. Cell Biol. 19(2), 176–184. Kaestner, K. H. (2005). The making of the liver: Developmental competence in foregut endoderm and induction of the hepatogenic program. Cell Cycle 4(9), 1146–1148. Kimelman, D., and Xu, W. (2006). beta-catenin destruction complex: Insights and questions from a structural perspective. Oncogene 25(57), 7482–7491. Kitisin, K., Saha, T., Blake, T., Golestaneh, N., Deng, M., Kim, C., et al. (2007). Tgf-Beta signaling in development. Sci. STKE 2007(399), cm1. Kuhl, M., Sheldahl, L. C., Park, M., Miller, J. R., and Moon, R. T. (2000). The Wnt/Ca2þ pathway: A new vertebrate Wnt signaling pathway takes shape. Trends Genet. 16(7), 279–283. Lemaigre, F., and Zaret, K. S. (2004). Liver development update: New embryo models, cell lineage control, and morphogenesis. Curr. Opin. Genet. Dev. 14(5), 582–590. Liu, P., Wakamiya, M., Shea, M. J., Albrecht, U., Behringer, R. R., and Bradley, A. (1999). Requirement for Wnt3 in vertebrate axis formation. Nat. Genet. 22(4), 361–365. Logan, C. Y., and Nusse, R. (2004). The Wnt signaling pathway in development and disease. Annu. Rev. Cell Dev. Biol. 20, 781–810. Lowe, L. A., Yamada, S., and Kuehn, M. R. (2001). Genetic dissection of nodal function in patterning the mouse embryo. Development 128(10), 1831–1843. Massague, J., Seoane, J., and Wotton, D. (2005). Smad transcription factors. Genes Dev. 19 (23), 2783–2810. Moon, R. T. (2005). Wnt/beta-catenin pathway. Sci. STKE 2005(271), cm1. Moon, R. T., Brown, J. D., and Torres, M. (1997). WNTs modulate cell fate and behavior during vertebrate development. Trends Genet. 13(4), 157–162. Moustakas, A. (2002). Smad signalling network. J. Cell. Sci. 115(Pt 17), 3355–3356. Nateri, A. S., Spencer-Dene, B., and Behrens, A. (2005). Interaction of phosphorylated c-Jun with TCF4 regulates intestinal cancer development. Nature 437(7056), 281–285.
216
Catherine Payne et al.
Ober, E. A., Verkade, H., Field, H. A., and Stainier, D. Y. (2006). Mesodermal Wnt2b signalling positively regulates liver specification. Nature 442(7103), 688–691. Rossi, J. M., Dunn, N. R., Hogan, B. L., and Zaret, K. S. (2001). Distinct mesodermal signals, including BMPs from the septum transversum mesenchyme, are required in combination for hepatogenesis from the endoderm. Genes Dev. 15(15), 1998–2009. Serls, A. E., Doherty, S., Parvatiyar, P., Wells, J. M., and Deutsch, G. H. (2005). Different thresholds of fibroblast growth factors pattern the ventral foregut into liver and lung. Development 132(1), 35–47. Sharma, R., Greenhough, S., Medine, C., and Hay, D. C. (2010). Three dimensional culture of human embryonic stem cell derived hepatic endoderm and its role in bioartificial liver construction. J. Biomed. Biotechnol. Vol. 2010. Article ID 236147. Strom, S. C., Fisher, R. A., Rubinstein, W. S., Barranger, J. A., Towbin, R. B., Charron, M., et al. (1997). Transplantation of human hepatocytes. Transplant. Proc. 29 (4), 2103–2106. Sullivan, G. J., Hay, D. C., Park, I. H., Fletcher, J., Hannoun, Z., Payne, C. M., et al. (2010). Generation of functional human hepatic endoderm from human induced pluripotent stem cells. Hepatology 51(1), 329–335. Tam, P. P., and Loebel, D. A. (2007). Gene function in mouse embryogenesis: Get set for gastrulation. Nat. Rev. Genet. 8(5), 368–381. Tam, P. P., Khoo, P. L., Wong, N., Tsang, T. E., and Behringer, R. R. (2004). Regionalization of cell fates and cell movement in the endoderm of the mouse gastrula and the impact of loss of Lhx1(Lim1) function. Dev. Biol. 274(1), 171–187. Topol, L., Jiang, X., Choi, H., Garrett-Beal, L., Carolan, P. J., and Yang, Y. (2003). Wnt-5a inhibits the canonical Wnt pathway by promoting GSK-3-independent beta-catenin degradation. J. Cell Biol. 162(5), 899–908. van Noort, M., and Clevers, H. (2002). TCF transcription factors, mediators of Wntsignaling in development and cancer. Dev. Biol. 244(1), 1–8. Westfall, T. A., Brimeyer, R., Twedt, J., Gladon, J., Olberding, A., Furutani-Seiki, M., et al. (2003). Wnt-5/pipetail functions in vertebrate axis formation as a negative regulator of Wnt/beta-catenin activity. J. Cell Biol. 162(5), 889–898. Wu, C. H., and Nusse, R. (2002). Ligand receptor interactions in the Wnt signaling pathway in Drosophila. J. Biol. Chem. 277(44), 41762–41769. Yamamoto, M., Meno, C., Sakai, Y., Shiratori, H., Mochida, K., Ikawa, Y., et al. (2001). The transcription factor FoxH1 (FAST) mediates Nodal signaling during anterior-posterior patterning and node formation in the mouse. Genes Dev. 15(10), 1242–1256. Zaret, K. S. (2001). Hepatocyte differentiation: From the endoderm and beyond. Curr. Opin. Genet. Dev. 11(5), 568–574. Zhao, R., and Duncan, S. A. (2005). Embryonic development of the liver. Hepatology 41(5), 956–967.
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Activin in Glucose Metabolism Osamu Hashimoto* and Masayuki Funaba† Contents I. Introduction II. Activin Activity in Insulin-Producing and Insulin-Sensitive Tissues A. Pancreas B. Liver C. Skeletal muscle D. Adipose tissues III. Activin in Adipose Tissue Inflammation IV. Conclusions and Future Directions Acknowledgments References
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Abstract Activins, members of the TGF-b family, are multifunctional growth and differentiation factors. Activins regulate glucose/energy metabolism by promoting the differentiation of insulin-producing and -responsive cells, and regulating function of the differentiated cells. In the pancreas, activins stimulate the differentiation of b cells and secretion of insulin, which enables the cells to respond to glucose uptake efficiently. By contrast, in the liver, skeletal muscle and white adipose tissue, activins exert negative regulation on organogenesis, which leads to impaired insulin sensitivity. Activins induce the phenotypic switch of macrophages from the M1 to M2 phenotypes, which reduces inflammation. Since adipose inflammation is closely associated with insulin resistance and the onset of type 2 diabetes, activins may improve insulin resistance through their anti-inflammatory activity. Because activins modulate events involved in insulin sensitivity in a tissue-dependent manner, the activities of activins should be locally regulated to improve whole-body insulin responsiveness. Thus, activins or activin inhibitors may be effective as therapeutic agents for metabolic syndrome. ß 2011 Elsevier Inc. * Laboratory of Experimental Animal Science, Faculty of Veterinary Medicine, Kitasato University, School of Veterinary Medicine, Towada, Aomori, Japan Division of Applied Biosciences, Kyoto University Graduate School of Agriculture, Kitashirakawa Oiwakecho, Kyoto, Japan
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Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00011-1
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I. Introduction Upon consumption of carbohydrates in mammals, digestion yields simple sugars such as glucose and fructose that are converted to pyruvate via glycolysis. In turn, pyruvate is oxidized to provide energy via the citrate cycle and the oxidative phosphorylation pathways. Thus, simple sugars act as a metabolic fuel. However, when excess carbohydrate is ingested, the surplus sugars are channeled into pathways for the synthesis of fatty acids after conversion to pyruvate, leading to fat accumulation and, ultimately, obesity. The blood glucose level is maintained within a narrow range principally by three hormones, insulin, glucagon, and adrenalin (epinephrine), in mammals. Glucagon and adrenalin elevate the blood glucose level by stimulating glycogenolysis, gluconeogenesis, and lipolysis. In contrast, insulin accelerates glucose uptake in the liver, muscle, and adipose tissues, and thus decreases the blood glucose level (Wasserman, 1995). Insulin is the only hormone that reduces the blood glucose level, and disturbances in insulin synthesis and responsiveness result in spontaneous damage of cellular and extracellular proteins in physiological systems through protein glycation (Ahmed and Thornalley, 2007). Thus, because of implications of glycation on health such as microvascular disease (nephropathy, retinopathy, neuropathy), the control of insulin production and its responses is particularly important for quality of life. During the evolutionary process, organisms, including mammals, have developed various mechanisms to defend against starvation, through efficient energy conversion during nutrient metabolism and by reducing energy expenditure during rest. However, organisms, including mammals, respond poorly to overfeeding. Therefore, long-term excess intake of energy leads to obesity. In fact, the prevalence of obesity has increased dramatically, particularly in economically advanced societies, in recent years (Flier, 2004). Obesity induces insulin resistance, which is associated with various diseases such as type 2 diabetes, coronary artery disease and hypertension, and the coexistence of these diseases is known as the metabolic syndrome (Kadowaki et al., 2006). Activins, members of the TGF-b family, regulate various biological processes. They were originally identified as stimulators of FSH secretion from pituitary cells and were isolated from the ovarian follicular fluid as disulfide-linked homo- or heterodimers of inhibin/activin bA or bB subunit, activin A (bAbA), activin AB (bAbB), and activin B (bBbB) (Wiater and Vale, 2008). In addition, the related genes were cloned in mammals as inhibin/activin bC and bE subunits (Deli et al., 2008) and seven other activin isoforms containing inhibin/activin bC and bE subunits are possible. Most of the studies have used activin A as the activin source. Similar to the
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other members of the TGF-b family, activins signal through two types of transmembrane serine/threonine kinase receptors, type I and type II receptors in target cells. ALK4 is the type I receptor, and ActRIIA and ActRIIB are the type II receptors (Tsuchida et al., 2009). In addition to ALK4, activins AB and B can transmit their signal via ALK7 as a type I receptor (Tsuchida et al., 2004). Upon activins binding, the type II receptor kinases transphosphorylate type I receptors, which then transmit the signals downstream by phosphorylating the carboxyl-terminal serines in their physiological substrates, Smad2 and Smad3. Once activated by phosphorylation, Smad2 and Smad3 form complexes with Smad4, and accumulate in the nucleus where they interact with the transcriptional regulators of the target genes (Tsuchida et al., 2009). Using molecular, cellular, and genetic approaches, activins have been shown to be associated with insulin production and responses. This article reviews the evidence underpinning the importance of activins in glucose/energy metabolism, particularly in insulin-producing and -responsive organs.
II. Activin Activity in Insulin-Producing and Insulin-Sensitive Tissues A. Pancreas Pancreatic islet cells are the main source of insulin and glucagon, which are produced by b cells and a cells, respectively. The secretion of hormones from pancreatic islets is mainly regulated by the glucose concentration. The growth and differentiation of hormone-producing cells and the secretion of hormones must be rigorously regulated to maintain glucose homeostasis. The gene expression of the inhibin/activin bB subunit and the canonical activin receptors (i.e., ALK4, ActRIIA, and ActRIIB) has been demonstrated in the pancreatic bud during gestation in mouse embryos (Dichmann et al., 2003), and the inhibin/activin bA subunit was localized in a, b, and d cells of the pancreatic islet (Ogawa et al., 1993; Yasuda et al., 1993a). Previous studies have demonstrated that activin B positively regulates pancreatic islet development and insulin secretion. Treatment with activin B stimulated the differentiation of fetal pancreatic endoderm into b cells and upregulated the expression of insulin (Hebrok et al., 1998). The activin B-induced differentiation resulted from downregulation of sonic hedgehog expression, which induced the homeobox transcription factor Pdx1 and thus determined b cell differentiation (Hebrok et al., 1998). Consistent with the effects of exogenous activin B, a marked reduction in the size and number of islets expressing Pdx1 and insulin was observed in ActRIIA and ActRIIB mutant mouse embryos (Kim et al., 2000). In addition, transgenic mice expressing dominant-negative ActRIIA and ActRIIB
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showed hypoplasia of pancreatic islets (Shiozaki et al., 1999; Yamaoka et al., 1998), and impaired glucose tolerance (Yamaoka et al., 1998). Furthermore, similar phenotypes were observed in adult ActRIIA- or ActRIIB-null mice (Goto et al., 2007; Kim et al., 2000). Activin-mediated development of pancreatic islets is also promoted during pancreatic regeneration in rats. The gene transcript levels of inhibin/activin bA and bB in the pancreatic duct were increased after streptozotocin injection, pancreatectomy, or pancreatic duct ligation, suggesting upregulation of activins A, AB, and B during neogenesis of b cells (Li et al., 2003; Zhang et al., 2002). Coadministration of activin A and b-cellulin, a member of the EGF family, reduced blood glucose levels and improved glucose tolerance in neonatal streptozotocin-treated rats, a model of b cell regeneration (Li et al., 2004). In addition, the administration of follistatin inhibited the differentiation of duct epithelial cells into b cells in another pancreatic regeneration model in which IFNg was expressed under the control of the insulin promoter in nonobese diabetic mice (Zhang et al., 2004). Since follistatin binds to and neutralizes the effects of activin (Sugino et al., 1997), the results are consistent with the positive effects of activins in pancreatic cell differentiation. The involvement of activins in the commitment of cells into pancreatic b cell lineage cells has been further verified in in vitro studies. Activin A induced the transdifferentiation of exocrine AR42J cells, a pancreatic cell line, into insulin-secreting endocrine cells (Mashima et al., 1996), and potentiated the differentiation of human fetal epithelial pancreatic cells into insulin-producing b cells (Demeterco et al., 2000). In a human embryonic stem (ES) cell line, treatment with activin A increased the population of Pdx1-positive pancreatic-endodermal cells, but not terminally differentiated insulin-producing cells ( Jiang et al., 2007a,b; Shim et al., 2007). In another ES cell line, activin B was reported to mediate cell differentiation into Pdx1-positive cells, whereas activin A promoted selfrenewal of the undifferentiated cells (Frandsen et al., 2007). These results suggest that activins potentially determine the cell fate of undifferentiated cells into pancreatic b cell lineage cells (Furth and Atala, 2009; Sulzbacher et al., 2009). Consistent with this, the addition of activin A to the culture medium was required for differentiation of induced pluripotent stem (iPS) cells into insulin-producing cells ( Jiang et al., 2007a; Tateishi et al., 2008). Activins have dual effects on insulin secretion; activins have potency for basal production and secretion of insulin, but they also prevent excess production and secretion in response to glucose stimulation. Activin B stimulated the secretion of insulin from MIN6 cells, a pancreatic b cell line (Tsuchida et al., 2004). Insulin secretion from cultured pancreatic islet cells was also increased in response to treatment with activin A (Florio et al., 2000; Totsuka et al., 1988; Verspohl et al., 1993). Activins AB and B, but not activin A, transduce the signal via ALK7, a type I receptor for the
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TGF-b family, and through ALK4 (Tsuchida et al., 2004; Zhang et al., 2006). Insulin gene transcription was accelerated by the activation of ALK7 in MIN6 cells (Watanabe et al., 2008). Collectively, these results suggest positive effects of activins on insulin production or secretion, or both. By contrast, activin B depressed glucose-stimulated Ca2þ influx, a trigger of insulin secretion in pancreatic islets (Bertolino et al., 2008). Mice with a null mutation of the inhibin/activin bB gene exhibited hyperinsulinemia (Bertolino et al., 2008). ALK7 knockout mice also showed elevated serum insulin levels, reduced insulin sensitivity and impaired glucose tolerance (Bertolino et al., 2008). In terms of its effects on a cells, activin potently inhibits the production and secretion of glucagon. Glucagon expression was transcriptionally inhibited by activin A treatment in InR1G9 and aTC1 cells, a cell lines (Mamin and Philippe, 2007). Activin A also inhibited glucagon secretion from rat pancreatic islets at a low glucose concentration (Verspohl et al., 1993).
B. Liver The liver plays a major role in carbohydrate metabolism. The metabolic fate of glucose is modulated depending on the energy status in the liver. Glycogenolysis and gluconeogenesis are stimulated in response to starvation, whereas feeding induces glycogen synthesis and lipogenesis. Insulin positively regulates energy storage processes. Activins modulate the growth and differentiation of hepatocytes, and glucose synthesis and release. Activin A was reported to inhibit mitogen-stimulated growth of parenchymal liver cells (Yasuda et al., 1993b). Inhibin/activin bA was expressed in parenchymal cells (De Bleser et al., 1997; Yasuda et al., 1993b), and treatment with follistatin enhanced mitogen-induced DNA synthesis in parenchymal cells (Yasuda et al., 1993b). Thus, activin A acts as an autocrine inhibitor of hepatocyte proliferation. During liver regeneration after partial hepatectomy, the expression of inhibin/activin bA was lower than that in sham-operated rats (Gold et al., 2005). However, the molecular basis for activin-induced growth arrest in hepatocytes remains poorly understood. In a human hepatoma cell line, HepG2, hypophosphorylation of retinoblastoma protein was involved in activin-mediated cell growth inhibition (Zauberman et al., 1997). In addition, activin treatment induced the expression of the cyclin-dependent kinase inhibitor p15 and transcription factor Sp1 (Ho et al., 2004). Activin A stimulated glycogenolysis in isolated rat hepatocytes in a dosedependent manner by increasing the cytoplasmic Ca2þ concentration (Mine et al., 1989), although it did not increase glucose production in perfused rat liver (Kojima et al., 1995). Activin also regulates gluconeogenesis, as treatment with activin enhanced the expression of glucose-6-phosphatase, a key
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gluconeogenesis enzyme that catalyzes the conversion of glucose6-phosphate to glucose, in HepG2 cells (Mukherjee et al., 2007). Unlike the inhibin/activin bA and bB subunits, inhibins bC and bE are predominantly expressed in the liver (Deli et al., 2008). Targeted disruption of inhibin/activin bC or bE, or both, showed no apparent effects on liver growth, differentiation, and regeneration, or on the serum biochemical profiles related to liver function (Lau et al., 2000). Thus, the biological activity of activins C and E has not been extensively evaluated, and their physiological roles are largely unknown. Hepatic inhibin/activin bC mRNA expression was increased in response to starvation in rats (Kogure et al., 1998), and the expression of inhibin/activin bE in the liver showed feeding-related diurnal changes in meal-fed rats, with a gradual increase before feeding and a rapid decrease after feeding (Rodgarkia-Dara et al., 2006). In addition, insulin transcriptionally upregulated inhibin/activin bE expression in liver cells (Hashimoto et al., 2009). Furthermore, transgenic mice overexpressing activin E showed higher insulin sensitivity and lower blood glucose levels than control mice, which was not due to modulation of pancreatic islet development (Hashimoto et al., 2006; Hashimoto, Unpublished data). Therefore, activin E and possibly activin C may be involved in the regulation of glucose/energy metabolism as a local or endocrine factor, or both.
C. Skeletal muscle Skeletal muscle is the largest tissue responsible for insulin-stimulated glucose disposal in the body. Thus, the preservation of muscle mass is important for the maintenance of the blood sugar level, and an increase in muscle mass improves insulin sensitivity. Activins modulate cell-fate determination to myoblast lineage cells and myotube differentiation of myoblasts, leading to the regulation of muscle mass. Overexpression of activin A in the tibialis anterior muscle caused muscle atrophy in rats (Gilson et al., 2009). Treatment with activin A inhibited multinucleated myotube formation in primary chicken muscle cell cultures in a dose-dependent manner, whereas follistatin enhanced myotube formation (Link and Nishi, 1997). Since inhibin/activin bA and its receptors are expressed in skeletal muscle (Hilde´n et al., 1994; Link and Nishi, 1997; Tuuri et al., 1994), activin A appears to be an autocrine negative regulator of myogenesis. This is supported by in vivo implant experiment in which beads coated with activin A were implanted into the chick limb bud and transiently downregulated the expression of Pax-3 and MyoD, transcription factors that are responsible for myoblast growth and differentiation (He et al., 2005). Although treatment with the activin isoforms A, B, and AB inhibited the differentiation of C2C12 myoblast cells into myotubes (Souza et al., 2008), the effective concentration was much higher than normal (4 mg/ml), because activin A was frequently used at concentrations of
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0.05–0.1 mg/ml in cell culture studies. In fact, treatment with 0.1 mg/ml activin A did not induce the phosphorylation of Smad2 in C2C12 myoblasts, whereas 5 ng/ml TGF-b1 caused significant Smad2 phosphorylation (Furutani et al., 2011). Thus, activin A may inhibit the commitment of mesenchymal stem cells to myoblast lineage cells, but not the differentiation of myoblasts into myotubes. The injection of soluble ActRIIB/Fc fusion protein increased hepatic insulin sensitivity in mice fed a high-fat diet, which was primarily due to muscle hypertrophy and secondarily to loss of fat deposits (Akpan et al., 2009). Consistently, transgenic expression of dominant-negative ActRIIB in skeletal muscle improved glucose tolerance in mice fed a standard diet or a high-fat diet, and resulted in resistance to diet-induced obesity (Guo et al., 2009). Myostatin, also referred to as GDF-8, is a potent negative regulator of myogenesis. For example, targeted disruption of the myostatin gene caused a dramatic and widespread increase in skeletal muscle mass in mice (McPherron et al., 1997). The myostatin signal transduction pathway partly overlaps with that of activins; myostatin forms a complex with ALK4 or ALK5 as the type I receptor and ActRIIB as the type II receptor, and transmitted its signal via Smad2 (Rebbapragada et al., 2003). Therefore, changes in insulin sensitivity caused by modulation of ActRIIB may be inhibited by myostatin activity. In fact, myostatin was strongly expressed in adult skeletal muscle (McPherron et al., 1997; Suryawan et al., 2006), whereas the concentrations of activin A in the skeletal muscle of rats increased from birth to 10 days old, and then markedly declined (Suryawan et al., 2006).
D. Adipose tissues There are two types of adipocytes; white adipose tissue (WAT) and brown adipose tissue (BAT). Adipocytes in WAT store energy as lipid, whereas those in BAT dissipate energy as heat by uncoupling oxidative phosphorylation (Cannon and Nedergaard, 2004). The functionally opposite adipocytes have long been proposed to share a common developmental origin, although a recent study revealed that brown adipocytes originated from a myoblast lineage (Seale et al., 2008). Accumulating evidence suggests the involvement of activins in adipocyte differentiation and in adipocyte function related to insulin resistance. 1. White adipose tissue In white adipocytes, insulin increases glucose uptake through membrane translocation of glucose transporter. Insulin also promotes lipogenesis and inhibits lipolysis. Adipokines, bioactive peptides such as adiponectin, leptin, and TNF-a, are produced and secreted from adipocytes via pathways associated with responses to insulin. Activins affect glucose/energy metabolism through the modulation of adipocyte differentiation and insulin sensitivity.
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Activin A was reported to inhibit adipocyte differentiation of 3T3-L1 preadipocytes (Hirai et al., 2005). Of note, activin A dose-dependently decreased the expression of peroxisome proliferative activated receptor-g2 (PPAR-g2), a master regulator for adipocyte differentiation. Adipocyte differentiation was also inhibited in activin A-treated ROB-C26 mesenchymal cells (Kawabata et al., 2007) and primary bovine stromal vascular cells (Hirai et al., 2007). Although, the molecular basis for activin A-induced inhibition is unclear, it has been suggested that binding of Smad3 and CCAAT/enhancer-binding protein (C/EBP)b or C/EBPd and the inhibition of PPAR-g2 transcription by repressing C/EBP transactivation function are involved in TGF-b-induced inhibition of adipocyte differentiation (Choy and Derynck, 2003). Activin B is a potential adipokine that regulates glucose/energy metabolism. Marked expression of inhibin/activin bB subunit was detected in human subcutaneous adipocytes (Sjo¨holm et al., 2006) and was positively related to the extent of obesity (Carlsson et al., 2009; Sjo¨holm et al., 2006). Increased expression of ALK7, a signal receptor for activins AB and B (Tsuchida et al., 2004; Zhang et al., 2006), was also detected in human adipose tissues (Carlsson et al., 2009), but the expression of ALK7 in subcutaneous adipose tissue was higher in lean than in obese subjects (Carlsson et al., 2009). In mice, the expression of inhibin/activin bB subunit was higher in epididymal adipose tissue than in other tissues, including subcutaneous adipose tissue (Hoggard et al., 2009). Furthermore, its expression in epididymal adipose tissue was higher in diabetic obese ob/ob mice than in lean mice. Interestingly, obese mice treated with leptin showed improved insulin resistance and decreased gene transcription, while insulin upregulated the expression of the inhibin/activin bB subunit in cultured adipocytes (Hoggard et al., 2009). Because of the significant role of the visceral fat depot in insulin resistance (Shibasaki et al., 2002), activin B may be involved in the improvement of insulin resistance. In fact, mice carrying an allele in which the mature region of the inhibin/activin bA subunit is replaced with the bB subunit had smaller fat pads and greater oxygen consumption compared with wild-type littermates (Li et al., 2009). The expression of genes related to the regulation of energy expenditure, including uncoupling protein (UCP), Pgc-1a, and FoxO1 in the liver, skeletal muscle and BAT were elevated in the mutant mice, explaining the leanness and higher metabolic rates of these mice (Li et al., 2009). Plasminogen activator inhibitor-1 (PAI-1) is also an adipokine, and increased expression of PAI-1 is associated with insulin resistance (Alessi et al., 2000). Since activated Smad3 and Smad4 confer transcription of the PAI-1 gene (Dennler et al., 1998), increased activity of activins may lead to insulin resistance.
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2. Brown adipose tissue As compared with the knowledge in WAT, the role of activins in BAT is limited. The roles of activins during differentiation of preadipocytes originating from BAT are broadly unknown. It is clear that the structurally related TGF-b1 acts as a mitogen and promotes differentiation of preadipocytes into brown adipocytes (Teruel et al., 1995, 1996), which was opposite to that in 3T3-L1 preadipocytes (Choy and Derynck, 2003). Upregulation of UCP1, the critical molecule for thermogenesis in BAT and the emergence of UCP1-positive adipocytes in WAT have been detected in transgenic mice overexpressing activin E (Hashimoto, Unpublished data). These may be related to greater insulin sensitivity in activin E-transgenic mice, as described above, and suggest the role of activin E for the prevention of insulin resistance.
III. Activin in Adipose Tissue Inflammation Macrophages are classified into two types, M1 and M2; M1 cells have proinflammatory activities, while M2 cells are anti-inflammatory (Mantovani et al., 2004). M1 macrophages infiltrate the adipose tissue of obese humans and mice, leading to adipose tissue inflammation. The inflammation induced by proinflammatory cytokines is closely associated with insulin resistance and the onset of type 2 diabetes (Hotamisligil and Erbay, 2008; Schenk et al., 2008). The administration of anti-inflammatory salicylates improved insulin responsiveness (Yuan et al., 2001), and inhibition of the NF-kB pathway, which is activated in response to inflammation, prevented the development of insulin resistance in mice (Cai et al., 2005). Treatment of macrophages with thiazolidinediones, insulin-sensitizing PPAR-g ligands, polarizes macrophages toward the M2 state. Therefore, M2 macrophages activated by PPAR-g might play role in anti-inflammation and enhance insulin sensitivity (Hevener et al., 2007). The expression of activin A was upregulated in peritoneal macrophages in response to their activation (Ogawa et al., 2000). In addition, treatment with activin A promoted the differentiation of peritoneal macrophages toward the M2 phenotype by inducing the expression of arginase-1, an inhibitor of iNOS, and decreased IFNg-induced expression of iNOS (Ogawa et al., 2006). It was also reported that activin A potentiated the inhibition of NO release in endotoxin-activated macrophages (Wang et al., 2008; Zhou et al., 2009). The role of activin A as an anti-inflammatory cytokine is also supported by the following results. Insulin stimulated the release of activin A in endotoxin-mediated inflammation, decreasing the production of the inflammatory mediators, TNF-a and IL-8, in macrophages (Cuschieri et al., 2008).
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Meanwhile, treatment with adiponectin induced the release of activin A from primary human monocytes (Weigert et al., 2009). Similar to macrophages in obese adipose tissues, inflammation induced by Kupffer cells may be linked to hepatic insulin resistance in obesity (Schenk et al., 2008). Inhibition of the NF-kB pathway in the liver improved hepatic insulin sensitivity (Schenk et al., 2008). Inflammation increased the expression of inhibin/activin bB and bE subunits in the liver ( Jones et al., 2007; O’Bryan et al., 2000). Thus, activin isoforms other than activin A are likely to prevent hepatic insulin resistance via their anti-inflammatory activities.
IV. Conclusions and Future Directions The regulatory expression of activins and the components involved in activin signaling in insulin-producing cells and in insulin-sensitive cells in response to changes in energy status undoubtedly indicates that the activin pathway is involved in fine-tuning of glucose/energy metabolism. As shown in Fig. 11.1, activins regulate glucose/energy metabolism in a complex manner. In the pancreas, activins stimulate the differentiation of b cells and insulin secretion, and thus facilitates efficient glucose uptake in other tissues. By contrast, in the liver, skeletal muscle and WAT, activins potentiated a decrease in organ mass, and thus impairs insulin sensitivity. However, activin increased the expression of a key enzyme in the gluconeogenesis pathway (Mukherjee et al., 2007), suggesting a dual role of activins in the liver. Activins induce a phenotypic switch from M1 to M2 in macrophages, which reduces inflammation. Since adipose tissue inflammation is closely associated with insulin resistance and the onset of type 2 diabetes, activins may improve insulin resistance through their antiinflammatory activities. Because activins modulate events involved in insulin sensitivity in a tissue-dependent and function-specific manner, their activities should be locally and finely regulated to improve whole-body insulin responsiveness. In terms of the biological activities and signal transduction of activins, the activin A isoform has been extensively studied. Although information on the other isoforms is limited, the functions of these other isoforms partly overlaps with those of activin A, but are distinct, as described above. In particular, the predominant localization in the liver and the feeding-related expression of activins C and E suggest their roles in homeostasis of the nutritional status through modulation of hormonal regulation. Unlike activin A, which potently regulates cell differentiation, activins C and E may protect against nutritional disturbances in differentiated cells. The absence
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Inflammation
WAT
Activin A
Islet regeneration
Activin A Activin B
Pancreas
Insulin sensitivity Differentiation Insulin secretion Activin B Insulin sensitivity
Insulin sensitivity Skeletal muscle Activin A
Insulin sensitivity
Liver
Activin A
Differentiation
BAT Differentiation
Activin E Activin C
Figure 11.1 Proposed roles of activins in glucose/energy metabolism. Activins regulate glucose/energy metabolism by modulating the differentiation and function of insulin-producing and insulin-sensitive cells. Solid lines indicate paracrine or autocrine activity. Dotted lines indicate endocrine activity. Arrows indicate activation and T indicates suppression.
of a prominent phenotype in mice with knockout of activins C and E (Lau et al., 2000) supports this possibility. Although the cross-talk between activin and insulin has not been extensively studied, a connection between insulin signaling and the TGF-b pathway, which shares the Smad-dependent pathway with activin signaling, has been demonstrated at various stages. Insulin signals via two major pathways, the mitogen-activated protein kinase (MAPK) and the phosphatidylinositol-3 kinase (PI3K) pathways (Le Roith and Zick, 2001). The metabolic responses to insulin are primarily mediated via the PI3K pathway; after the p85/p110 complex of PI3K associates with the insulin receptor substrate molecule, PI3K increases the production of phosphatidylinositol 3,4,5-phosphate (PIP3). In turn, PIP3 binds to PI3K-dependent kinase (PDK)-1 and Akt. This activates PDK1, which phosphorylates and activates Akt. The activated Akt phosphorylates proteins including FoxO, Forkhead transcription factors and mTOR. PTEN dephosphorylates PIP3 and reverses the action of PI3K (Le Roith and Zick, 2001; Maehama et al., 2001). The MAPKs, ERK1/2, p38 MAPK and JNK, are activated by activin and TGF-b, and the activated MAPKs phosphorylate and modulate Smad2- and
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Smad3-mediated transcriptional activation in a cell type-specific manner (Tsuchida et al., 2009; Zhang, 2008). Glucose-induced hypertrophy of fibroblasts and epithelial cells is due to TGF-b-dependent activation of the Akt/mTOR pathway (Wu and Derynck, 2009). Akt was reported to modulate Smad3-mediated transcription through the direct interaction and sequestration of unphosphorylated Smad3 (Conery et al., 2004). Smad2 and Smad3 interact with FoxO to induce transcription of the p21Cip1 gene (Seoane et al., 2004). Another Forkhead transcription factor FoxC2, which counteracts insulin resistance in adipose tissues (Cederberg et al., 2001), was reported to associate with Smad3 and Smad4 and cooperatively activated PAI-1 gene transcription (Fujita et al., 2006). Meanwhile, the expression of PTEN was downregulated by TGF-b (Yang et al., 2009). However, these data remain limited, and the significance of the functional and physical interactions with the insulin-mediated cell responses are still unknown. The molecular mechanisms by which activin regulates insulin sensitivity should also be clarified in future studies. Such knowledge should help identify targets for the prevention and therapy of insulin resistance and type 2 diabetes.
ACKNOWLEDGMENTS The authors thank Dr. K. Sekiyama for helpful discussions. This work was supported by KAKENHI from the Japanese Society for the Promotion of Science (No. 21580370).
REFERENCES Ahmed, N., and Thornalley, P. J. (2007). Advanced glycation endproducts: What is their relevance to diabetic complications? Diabetes Obes. Metab. 9, 233–245. Akpan, I., Goncalves, M. D., Dhir, R., Yin, X., Pistilli, E. E., Bogdanovich, S., Khurana, T. S., Ucran, J., Lachey, J., and Ahima, R. S. (2009). The effects of a soluble activin type IIB receptor on obesity and insulin sensitivity. Int. J. Obes. 33, 1265–1273. Alessi, M. C., Bastelica, D., Morange, P., Berthet, B., Leduc, I., Verdier, M., Geel, O., and Juhan-Vague, I. (2000). Plasminogen activator inhibitor 1, transforming growth factor-b 1, and BMI are closely associated in human adipose tissue during morbid obesity. Diabetes 49, 1374–1380. Bertolino, P., Holmberg, R., Reissmann, E., Andersson, O., Berggren, P. O., and Iba´n˜ez, C. F. (2008). Activin B receptor ALK7 is a negative regulator of pancreatic b-cell function. Proc. Natl. Acad. Sci. USA 105, 7246–7251. Cai, D., Yuan, M., Frantz, D. F., Melendez, P. A., Hansen, L., Lee, J., and Shoelson, S. E. (2005). Local and systemic insulin resistance resulting from hepatic activation of IKK-b and NF-kB. Nat. Med. 11, 183–190. Cannon, B., and Nedergaard, J. (2004). Brown adipose tissue: Function and physiological significance. Physiol. Rev. 84, 277–359. Carlsson, L. M., Jacobson, P., Walley, A., Froguel, P., Sjo¨stro¨m, L., Svensson, P. A., and Sjo¨holm, K. (2009). ALK7 expression is specific for adipose tissue, reduced in obesity and
Role of Activins in Glucose/Energy Metabolism
229
correlates to factors implicated in metabolic disease. Biochem. Biophys. Res. Commun. 382, 309–314. Cederberg, A., Gr nning, L. M., Ahre´n, B., Taske´n, K., Carlsson, P., and Enerba¨ck, S. (2001). FOXC2 is a winged helix gene that counteracts obesity, hypertriglyceridemia, and diet-induced insulin resistance. Cell 106, 563–573. Choy, L., and Derynck, R. (2003). Transforming growth factor-b inhibits adipocyte differentiation by Smad3 interacting with CCAAT/enhancer-binding protein (C/EBP) and repressing C/EBP transactivation function. J. Biol. Chem. 278, 9609–9619. Conery, A. R., Cao, Y., Thompson, E. A., Townsend, C. M., Jr., Ko, T. C., and Luo, K. (2004). Akt interacts directly with Smad3 to regulate the sensitivity to TGF-b induced apoptosis. Nat. Cell Biol. 6, 366–372. Cuschieri, J., Bulger, E., Grinsell, R., Garcia, I., and Maier, R. V. (2008). Insulin regulates macrophage activation through activin A. Shock 29, 285–290. De Bleser, P. J., Niki, T., Xu, G., Rogiers, V., and Geerts, A. (1997). Localization and cellular sources of activins in normal and fibrotic rat liver. Hepatology 26, 905–912. Deli, A., Kreidl, E., Santifaller, S., Trotter, B., Seir, K., Berger, W., Schulte-Hermann, R., Rodgarkia-Dara, C., and Grusch, M. (2008). Activins and activin antagonists in hepatocellular carcinoma. World J. Gastroenterol. 14, 1699–1709. Demeterco, C., Beattie, G. M., Dib, S. A., Lopez, A. D., and Hayek, A. (2000). A role for activin A and betacellulin in human fetal pancreatic cell differentiation and growth. J. Clin. Endocrinol. Metab. 85, 3892–3897. Dennler, S., Itoh, S., Vivien, D., ten Dijke, P., Huet, S., and Gauthier, J. M. (1998). Direct binding of Smad3 and Smad4 to critical TGF b-inducible elements in the promoter of human plasminogen activator inhibitor-type 1 gene. EMBO J. 17, 3091–3100. Dichmann, D. S., Miller, C. P., Jensen, J., Scott Heller, R., and Serup, P. (2003). Expression and misexpression of members of the FGF and TGFb families of growth factors in the developing mouse pancreas. Dev. Dyn. 226, 663–674. Flier, J. S. (2004). Obesity wars: Molecular progress confronts an expanding epidemic. Cell 116, 337–350. Florio, P., Luisi, S., Marchetti, P., Lupi, R., Cobellis, L., Falaschi, C., Sugino, H., Navalesi, R., Genazzani, A. R., and Petraglia, F. (2000). Activin A stimulates insulin secretion in cultured human pancreatic islets. J. Endocrinol. Invest. 23, 231–234. Frandsen, U., Porneki, A. D., Floridon, C., Abdallah, B. M., and Kassem, M. (2007). Activin B mediated induction of Pdx1 in human embryonic stem cell derived embryoid bodies. Biochem. Biophys. Res. Commun. 362, 568–574. Fujita, H., Kang, M., Eren, M., Gleaves, L. A., Vaughan, D. E., and Kume, T. (2006). Foxc2 is a common mediator of insulin and transforming growth factor b signaling to regulate plasminogen activator inhibitor type I gene expression. Circ. Res. 98, 626–634. Furth, M. E., and Atala, A. (2009). Stem cell sources to treat diabetes. J. Cell. Biochem. 106, 507–511. Furutani, Y., Umemoto, T., Murakami, M., Matsui, T., and Funaba, M. (2011). Role of endogenous TGF-b family in myogenic differentiation of C2C12 cells. J. Cell. Biochem. In press. Gilson, H., Schakman, O., Kalista, S., Lause, P., Tsuchida, K., and Thissen, J. P. (2009). Follistatin induces muscle hypertrophy through satellite cell proliferation and inhibition of both myostatin and activin. Am. J. Physiol. Endocrinol. Metab. 297, E157–E164. Gold, E. J., Zhang, X., Wheatley, A. M., Mellor, S. L., Cranfield, M., Risbridger, G. P., Groome, N. P., and Fleming, J. S. (2005). bA- and bC-activin, follistatin, activin receptor mRNA and bC-activin peptide expression during rat liver regeneration. J. Mol. Endocrinol. 34, 505–515. Goto, Y., Nomura, M., Tanaka, K., Kondo, A., Morinaga, H., Okabe, T., Yanase, T., Nawata, H., Takayanagi, R., and Li, E. (2007). Genetic interactions between activin type
230
Osamu Hashimoto and Masayuki Funaba
IIB receptor and Smad2 genes in asymmetrical patterning of the thoracic organs and the development of pancreas islets. Dev. Dyn. 236, 2865–2874. Guo, T., Jou, W., Chanturiya, T., Portas, J., Gavrilova, O., and McPherron, A. C. (2009). Myostatin inhibition in muscle, but not adipose tissue, decreases fat mass and improves insulin sensitivity. PLoS ONE 4, e4937. Hashimoto, O., Ushiro, Y., Sekiyama, K., Yamaguchi, O., Yoshioka, K., Mutoh, K., and Hasegawa, Y. (2006). Impaired growth of pancreatic exocrine cells in transgenic mice expressing human activin bE subunit. Biochem. Biophys. Res. Commun. 341, 416–424. Hashimoto, O., Sekiyama, K., Matsuo, T., and Hasegawa, Y. (2009). The role of activin E in glucose metabolism: Transcriptional regulation of the inhibin/activin bE subunit gene in the liver. Life Sci. 85, 534–540. He, L., Vichev, K., Huang, R., Christ, B., Patel, K., and Amthor, H. (2005). Activin A inhibits formation of skeletal muscle during chick development. Anat. Embryol. 209, 401–407. Hebrok, M., Kim, S. K., and Melton, D. A. (1998). Notochord repression of endodermal Sonic hedgehog permits pancreas development. Genes Dev. 12, 1705–1713. Hevener, A. L., Olefsky, J. M., Reichart, D., Nguyen, M. T., Bandyopadyhay, G., Leung, H. Y., Watt, M. J., Benner, C., Febbraio, M. A., Nguyen, A. K., Folian, B., Subramaniam, S., et al. (2007). Macrophage PPAR-g is required for normal skeletal muscle and hepatic insulin sensitivity and full antidiabetic effects of thiazolidinediones. J. Clin. Invest. 117, 1658–1669. Hilde´n, K., Tuuri, T., Era¨maa, M., and Ritvos, O. (1994). Expression of type II activin receptor genes during differentiation of human K562 cells and cDNA cloning of the human type IIB activin receptor. Blood 83, 2163–2170. Hirai, S., Yamanaka, M., Kawachi, H., Matsui, T., and Yano, H. (2005). Activin A inhibits differentiation of 3T3-L1 preadipocyte. Mol. Cell. Endocrinol. 232, 21–26. Hirai, S., Matsumoto, H., Moriya, N. H., Kawachi, H., and Yano, H. (2007). Follistatin rescues the inhibitory effect of activin A on the differentiation of bovine preadipocyte. Domest. Anim. Endocrinol. 33, 269–280. Ho, J., de Guise, C., Kim, C., Lemay, S., Wang, X. F., and Lebrun, J. J. (2004). Activin induces hepatocyte cell growth arrest through induction of the cyclin-dependent kinase inhibitor p15INK4B and Sp1. Cell. Signal. 16, 693–701. Hoggard, N., Cruickshank, M., Moar, K. M., Barrett, P., Bashir, S., and Miller, J. D. (2009). Inhibin bB expression in murine adipose tissue and its regulation by leptin, insulin and dexamethasone. J. Mol. Endocrinol. 43, 171–177. Hotamisligil, G. S., and Erbay, E. (2008). Nutrient sensing and inflammation in metabolic diseases. Nat. Rev. Immunol. 8, 923–934. Jiang, J., Au, M., Lu, K., Eshpeter, A., Korbutt, G., Fisk, G., and Majumdar, A. S. (2007a). Generation of insulin-producing islet-like clusters from human embryonic stem cells. Stem Cells 25, 1940–1953. Jiang, W., Shi, Y., Zhao, D., Chen, S., Yong, J., Zhang, J., Qing, T., Sun, X., Zhang, P., Ding, M., Li, D., and Deng, H. (2007b). In vitro derivation of functional insulinproducing cells from human embryonic stem cells. Cell Res. 17, 333–344. Jones, K. L., Mansell, A., Patella, S., Scott, B. J., Hedger, M. P., de Kretser, D. M., and Phillips, D. J. (2007). Activin A is a critical component of the inflammatory response, and its binding protein, follistatin, reduces mortality in endotoxemia. Proc. Natl. Acad. Sci. USA 104, 16239–16244. Kadowaki, T., Yamauchi, T., Kubota, N., Hara, K., Ueki, K., and Tobe, K. (2006). Adiponectin and adiponectin receptors in insulin resistance, diabetes, and the metabolic syndrome. J. Clin. Invest. 116, 1784–1792.
Role of Activins in Glucose/Energy Metabolism
231
Kawabata, N., Kamiya, N., Suzuki, N., Matsumoto, M., and Takagi, M. (2007). Changes in extracellular activin A: Follistatin ratio during differentiation of a mesenchymal progenitor cell line, ROB-C26 into osteoblasts and adipocytes. Life Sci. 81, 8–18. Kim, S. K., Hebrok, M., Li, E., Oh, S. P., Schrewe, H., Harmon, E. B., Lee, J. S., and Melton, D. A. (2000). Activin receptor patterning of foregut organogenesis. Genes Dev. 14, 1866–1871. Kogure, K., Zhang, Y. Q., Shibata, H., and Kojima, I. (1998). Immediate onset of DNA synthesis in remnant rat liver after 90% hepatectomy by an administration of follistatin. J. Hepatol. 29, 977–984. Kojima, I., Nobusawa, R., Zhang, Y. Q., Sekine, N., Mine, T., and Shibata, H. (1995). Attenuation of glycogenolytic action of activin A in intact rat liver. Am. J. Physiol. 269, E846–E851. Lau, A. L., Kumar, T. R., Nishimori, K., Bonadio, J., and Matzuk, M. M. (2000). Activin bC and bE genes are not essential for mouse liver growth, differentiation, and regeneration. Mol. Cell. Biol. 20, 6127–6137. Le Roith, D., and Zick, Y. (2001). Recent advances in our understanding of insulin action and insulin resistance. Diabetes Care 24, 588–597. Li, M., Miyagawa, J., Moriwaki, M., Yuan, M., Yang, Q., Kozawa, J., Yamamoto, K., Imagawa, A., Iwahashi, H., Tochino, Y., Yamagata, K., and Matsuzawa, Y. (2003). Analysis of expression profiles of islet-associated transcription and growth factors during b-cell neogenesis from duct cells in partially duct-ligated mice. Pancreas 27, 345–355. Li, L., Yi, Z., Seno, M., and Kojima, I. (2004). Activin A and betacellulin: Effect on regeneration of pancreatic b-cells in neonatal streptozotocin-treated rats. Diabetes 53, 608–615. Li, L., Shen, J. J., Bournat, J. C., Huang, L., Chattopadhyay, A., Li, Z., Shaw, C., Graham, B. H., and Brown, C. W. (2009). Activin signaling: Effects on body composition and mitochondrial energy metabolism. Endocrinology 150, 3521–3529. Link, B. A., and Nishi, R. (1997). Opposing effects of activin A and follistatin on developing skeletal muscle cells. Exp. Cell Res. 233, 350–362. Maehama, T., Taylor, G. S., and Dixon, J. E. (2001). PTEN and myotubularin: Novel phosphoinositide phosphatases. Annu. Rev. Biochem. 70, 247–279. Mamin, A., and Philippe, J. (2007). Activin A decreases glucagon and arx gene expression in a-cell lines. Mol. Endocrinol. 21, 259–273. Mantovani, A., Sica, A., Sozzani, S., Allavena, P., Vecchi, A., and Locati, M. (2004). The chemokine system in diverse forms of macrophage activation and polarization. Trends Immunol. 25, 677–686. Mashima, H., Shibata, H., Mine, T., and Kojima, I. (1996). Formation of insulin-producing cells from pancreatic acinar AR42J cells by hepatocyte growth factor. Endocrinology 137, 3969–3976. McPherron, A. C., Lawler, A. M., and Lee, S. J. (1997). Regulation of skeletal muscle mass in mice by a new TGF-b family member. Nature 387, 83–90. Mine, T., Kojima, I., and Ogata, E. (1989). Stimulation of glucose production by activin-A in isolated rat hepatocytes. Endocrinology 125, 586–591. Mukherjee, A., Sidis, Y., Mahan, A., Raher, M. J., Xia, Y., Rosen, E. D., Bloch, K. D., Thomas, M. K., and Schneyer, A. L. (2007). FSTL3 deletion reveals roles for TGF-b family ligands in glucose and fat homeostasis in adults. Proc. Natl. Acad. Sci. USA 104, 1348–1353. O’Bryan, M. K., Sebire, K. L., Gerdprasert, O., Hedger, M. P., Hearn, M. T., and de Kretser, D. M. (2000). Cloning and regulation of the rat activin bE subunit. J. Mol. Endocrinol. 24, 409–418.
232
Osamu Hashimoto and Masayuki Funaba
Ogawa, K., Abe, K., Kurosawa, N., Kurohmaru, M., Sugino, H., Takahashi, M., and Hayashi, Y. (1993). Expression of a bA and bB subunits of inhibin or activin and follistatin in rat pancreatic islets. FEBS Lett. 319, 217–220. Ogawa, K., Funaba, M., Mathews, L. S., and Mizutani, T. (2000). Activin A stimulates type IV collagenase (matrix metalloproteinase-2) production in mouse peritoneal macrophages. J. Immunol. 165, 2997–3003. Ogawa, K., Funaba, M., Chen, Y., and Tsujimoto, M. (2006). Activin A functions as a Th2 cytokine in the promotion of the alternative activation of macrophages. J. Immunol. 177, 6787–6794. Rebbapragada, A., Benchabane, H., Wrana, J. L., Celeste, A. J., and Attisano, L. (2003). Myostatin signals through a transforming growth factor b-like signaling pathway to block adipogenesis. Mol. Cell. Biol. 23, 7230–7242. Rodgarkia-Dara, C., Vejda, S., Erlach, N., Losert, A., Brusch, W., Berger, W., SchulteHermann, R., and Grusch, M. (2006). The activin axis in liver biology and disease. Mutat. Res. 613, 123–137. Schenk, S., Saberi, M., and Olefsky, J. M. (2008). Insulin sensitivity: Modulation by nutrients and inflammation. J. Clin. Invest. 118, 2992–3002. Seale, P., Bjork, B., Yang, W., Kajimura, S., Chin, S., Kuang, S., Scime`, A., Devarakonda, S., Conroe, H. M., Erdjument-Bromage, H., Tempst, P., Rudnicki, M. A., et al. (2008). PRDM16 controls a brown fat/skeletal muscle switch. Nature 454, 961–967. Seoane, J., Le, H. V., Shen, L., Anderson, S. A., and Massague´, J. (2004). Integration of Smad and forkhead pathways in the control of neuroepithelial and glioblastoma cell proliferation. Cell 117, 211–223. Shibasaki, M., Takahashi, K., Itou, T., Miyazawa, S., Ito, M., Kobayashi, J., Bujo, H., and Saito, Y. (2002). Alterations of insulin sensitivity by the implantation of 3T3-L1 cells in nude mice. A role for TNF-a? Diabetologia 45, 518–526. Shim, J. H., Kim, S. E., Woo, D. H., Kim, S. K., Oh, C. H., McKay, R., and Kim, J. H. (2007). Directed differentiation of human embryonic stem cells towards a pancreatic cell fate. Diabetologia 50, 1228–1238. Shiozaki, S., Tajima, T., Zhang, Y. Q., Furukawa, M., Nakazato, Y., and Kojima, I. (1999). Impaired differentiation of endocrine and exocrine cells of the pancreas in transgenic mouse expressing the truncated type II activin receptor. Biochim. Biophys. Acta 1450, 1–11. Sjo¨holm, K., Palming, J., Lystig, T. C., Jennische, E., Woodruff, T. K., Carlsson, B., and Carlsson, L. M. (2006). The expression of inhibin bB is high in human adipocytes, reduced by weight loss, and correlates to factors implicated in metabolic disease. Biochem. Biophys. Res. Commun. 344, 1308–1314. Souza, T. A., Chen, X., Guo, Y., Sava, P., Zhang, J., Hill, J. J., Yaworsky, P. J., and Qiu, Y. (2008). Proteomic identification and functional validation of activins and bone morphogenetic protein 11 as candidate novel muscle mass regulators. Mol. Endocrinol. 22, 2689–2702. Sugino, H., Sugino, K., Hashimoto, O., Shoji, H., and Nakamura, T. (1997). Follistatin and its role as an activin-binding protein. J. Med. Invest. 44, 1–14. Sulzbacher, S., Schroeder, I. S., Truong, T. T., and Wobus, A. M. (2009). Activin Ainduced differentiation of embryonic stem cells into endoderm and pancreatic progenitors-the influence of differentiation factors and culture conditions. Stem Cell Rev. 5, 159–173. Suryawan, A., Frank, J. W., Nguyen, H. V., and Davis, T. A. (2006). Expression of the TGF-b family of ligands is developmentally regulated in skeletal muscle of neonatal rats. Pediatr. Res. 59, 175–179.
Role of Activins in Glucose/Energy Metabolism
233
Tateishi, K., He, J., Taranova, O., Liang, G., D’Alessio, A. C., and Zhang, Y. (2008). Generation of insulin-secreting islet-like clusters from human skin fibroblasts. J. Biol. Chem. 283, 31601–31607. Teruel, T., Valverde, A. M., Benito, M., and Lorenzo, M. (1995). Transforming growth factor b1 induces differentiation-specific gene expression in fetal rat brown adipocytes. FEBS Lett. 364, 193–197. Teruel, T., Valverde, A. M., Benito, M., and Lorenzo, M. (1996). Transforming growth factor b1 induces mitogenesis in fetal rat brown adipocytes. J. Cell. Physiol. 166, 577–584. Totsuka, Y., Tabuchi, M., Kojima, I., Shibai, H., and Ogata, E. (1988). A novel action of activin A: Stimulation of insulin secretion in rat pancreatic islets. Biochem. Biophys. Res. Commun. 156, 335–339. Tsuchida, K., Nakatani, M., Yamakawa, N., Hashimoto, O., Hasegawa, Y., and Sugino, H. (2004). Activin isoforms signal through type I receptor serine/threonine kinase ALK7. Mol. Cell. Endocrinol. 220, 59–65. Tsuchida, K., Nakatani, M., Hitachi, K., Uezumi, A., Sunada, Y., Ageta, H., and Inokuchi, K. (2009). Activin signaling as an emerging target for therapeutic interventions. Cell Commun. Signal. 7, 15. Tuuri, T., Era¨maa, M., Hilde´n, K., and Ritvos, O. (1994). The tissue distribution of activin bA- and bB-subunit and follistatin messenger ribonucleic acids suggests multiple sites of action for the activin-follistatin system during human development. J. Clin. Endocrinol. Metab. 78, 1521–1524. Verspohl, E. J., Ammon, H. P., and Wahl, M. A. (1993). Activin A: Its effects on rat pancreatic islets and the mechanism of action involved. Life Sci. 53, 1069–1078. Wang, S. Y., Tai, G. X., Zhang, P. Y., Mu, D. P., Zhang, X. J., and Liu, Z. H. (2008). Inhibitory effect of activin A on activation of lipopolysaccharide-stimulated mouse macrophage RAW264.7 cells. Cytokine 42, 85–91. Wasserman, D. H. (1995). Regulation of glucose fluxes during exercise in the postabsorptive state. Annu. Rev. Physiol. 57, 191–218. Watanabe, R., Shen, Z. P., Tsuda, K., and Yamada, Y. (2008). Insulin gene is a target in activin receptor-like kinase 7 signaling pathway in pancreatic b-cells. Biochem. Biophys. Res. Commun. 377, 867–872. Weigert, J., Neumeier, M., Wanninger, J., Schober, F., Sporrer, D., Weber, M., Schramm, A., Wurm, S., Sto¨gbauer, F., Filarsky, M., Scha¨ffler, A., Aslanidis, C., et al. (2009). Adiponectin upregulates monocytic activin A but systemic levels are not altered in obesity or type 2 diabetes. Cytokine 45, 86–91. Wiater, E., and Vale, W. (2008). Activins and inhibins. In “The TGF-b Family,” (R. Derynck and K. Miyazono, Eds.), pp. 79–120. Cold Spring Harbor Laboratory Press, New York. Wu, L., and Derynck, R. (2009). Essential role of TGF-b signaling in glucose-induced cell hypertrophy. Dev. Cell 17, 35–48. Yamaoka, T., Idehara, C., Yano, M., Matsushita, T., Yamada, T., Ii, S., Moritani, M., Hata, J., Sugino, H., Noji, S., and Itakura, M. (1998). Hypoplasia of pancreatic islets in transgenic mice expressing activin receptor mutants. J. Clin. Invest. 102, 294–301. Yang, Y., Zhou, F., Fang, Z., Wang, L., Li, Z., Sun, L., Wang, C., Yao, W., Cai, X., Jin, J., and Zha, X. (2009). Post-transcriptional and post-translational regulation of PTEN by transforming growth factor-b1. J. Cell. Biochem. 106, 1102–1112. Yasuda, H., Inoue, K., Shibata, H., Takeuchi, T., Eto, Y., Hasegawa, Y., Sekine, N., Totsuka, Y., Mine, T., and Ogata, E. (1993a). Existence of activin-A in A- and D-cells of rat pancreatic islet. Endocrinology 133, 624–630.
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Yasuda, H., Mine, T., Shibata, H., Eto, Y., Hasegawa, Y., Takeuchi, T., Asano, S., and Kojima, I. (1993b). Activin A: An autocrine inhibitor of initiation of DNA synthesis in rat hepatocytes. J. Clin. Invest. 92, 1491–1496. Yuan, M., Konstantopoulos, N., Lee, J., Hansen, L., Li, Z. W., Karin, M., and Shoelson, S. E. (2001). Reversal of obesity- and diet-induced insulin resistance with salicylates or targeted disruption of Ikkb. Science 293, 1673–1677. Zauberman, A., Oren, M., and Zipori, D. (1997). Involvement of p21WAF1/Cip1, CDK4 and Rb in activin A mediated signaling leading to hepatoma cell growth inhibition. Oncogene 15, 1705–1711. Zhang, Y. E. (2008). Non-Smad TGF-b signaling pathways. In “The TGF-b Family,” (R. Derynck and K. Miyazono, Eds.), pp. 419–437. Cold Spring Harbor Laboratory Press, New York. Zhang, Y. Q., Zhang, H., Maeshima, A., Kurihara, H., Miyagawa, J., Takeuchi, T., and Kojima, I. (2002). Up-regulation of the expression of activins in the pancreatic duct by reduction of the b-cell mass. Endocrinology 143, 3540–3547. Zhang, Y.-Q., Cleary, M. M., Si, Y., Liu, G., Eto, Y., Kritzik, M., Dabernat, S., Kayali, A. G., and Sarvetnick, N. (2004). Inhibition of activin signaling induces pancreatic epithelial cell expansion and diminishes terminal differentiation of pancreatic b-cells. Diabetes 53, 2024–2033. Zhang, N., Kumar, M., Xu, G., Ju, W., Yoon, T., Xu, E., Huang, X., Gaisano, H., Peng, C., and Wang, Q. (2006). Activin receptor-like kinase 7 induces apoptosis of pancreatic beta cells and beta cell lines. Diabetologia 49, 506–518. Zhou, J., Tai, G., Liu, H., Ge, J., Feng, Y., Chen, F., Yu, F., and Liu, Z. (2009). Activin A down-regulates the phagocytosis of lipopolysaccharide-activated mouse peritoneal macrophages in vitro and in vivo. Cell. Immunol. 255, 69–75.
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Activin in Humoral Immune Responses Kenji Ogawa* and Masayuki Funaba† Contents I. Introduction II. Activin A in Th2 Cells A. Expression of activin A exclusively in T helper cells but not in regulatory T cells B. Preferential production of activin A in Th2 cells C. Transactivation of the activin bA promoter by NFAT and c-Maf in Th2 cells III. Activin A in B Cells A. Production of activin A in activated B cells B. Involvement of activin A in immunoglobulin production in B cells IV. Activin A in Macrophages A. Activation-induced expression of activin A in macrophages B. Production of type IV collagenase in macrophages stimulated by activin A C. Alternative activation of macrophages induced by activin A V. Activin A in Mast Cells A. Upregulation of activin A expression in activated mast cells B. Modulation of mast cell function by activin A VI. Conclusions and Future Directions References
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Abstract The synthesis and secretion of activin A are stimulated in immune cells, including Th2 cells, B cells, peritoneal macrophages, dendritic cells, and mast cells, in response to their activation. The increased expression of activin A is regulated at the level of transcription. Activin A increases antigen-specific IgE production, promotes the differentiation of macrophages into the M2 phenotype, and stimulates the maturation of mast cells in an autocrine/paracrine fashion. * Molecular Ligand Discovery Research Team, Chemical Genomics Research Group, ASI, RIKEN, Hirosawa, Wako, Saitama, Japan Division of Applied Biosciences, Kyoto University Graduate School of Agriculture, Kitashirakawa Oiwakecho, Kyoto, Japan
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Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00012-3
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2011 Elsevier Inc. All rights reserved.
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The roles of activin A in Th2 cells, B cells, and peritoneal macrophages are distinct from those of TGF-b, which acts primarily as an immune suppressor. In contrast, the activities of activin A and TGF-b are similar in mast cells. These findings suggest the discrete regulation of immune responses by activin and TGF-b, two structurally related growth/differentiation factors. ß 2011 Elsevier Inc.
I. Introduction Activins, which are homo- or heterodimers of activin bA and bB subunits and members of the transforming growth factor (TGF)-b superfamily, are local regulators of cell growth and differentiation. The diverse physiological functions of activins in neural and endocrine tissues include stimulation of follicle-stimulating hormone (Vale et al., 1990; Ying, 1988) and prolactin (Murata and Ying, 1991) secretion from pituitary cells, inhibition of pituitary growth hormone secretion (Bilezikjian et al., 1990), regulation of nerve cell survival and differentiation (Hashimoto et al., 1990; Schubert et al., 1990) and neural differentiation in Xenopus embryos (Hemmati-Brivanlou and Melton, 1994; Hemmati-Brivanlou et al., 1994), stimulation of insulin secretion from pancreatic islets (Totsuka et al., 1988), and stimulation of steroidogenesis in gonadal cells (Li et al., 1992; Mauduit et al., 1991; Miro et al., 1991). In addition to their roles in neural and endocrine tissues, the expression and function of activins in the immune system have also been studied, although much less is known about this than about the roles of the structurally related TGF-b (Letterio, 2000). Activin-deficient mice are not viable and die within 24 h of birth, primarily due to craniofacial defects and the lack of a lower incisor (Matzuk et al., 1995a,b). The in vivo expression and function of activins in the immune system remain largely to be determined. In contrast, the immune suppressive function of TGF-b has been elegantly illustrated by in vivo studies. Targeted disruption of TGF-b1 in mice resulted in severe multifocal inflammation, indicating that it plays a role in immune suppression (Shull et al., 1992). TGF-b inhibits activated macrophage tumoricidal activity and cytokine production (Chen et al., 2008). Further, it inhibits lymphocyte function by inhibiting proliferation and differentiation into effector cells, and indeed induces apoptosis in B and T cells (Chen et al., 2008; Dennler et al., 2002). In most studies, activins have shown overlapping biological activities with TGF-b, partly as a result of the fact that activins and TGF-b molecules activate the same signaling intermediates: Smad2 and/or Smad3 (Lebrun et al., 1997; Massague´ and Chen, 2000). Thus, it was expected that activin A may also exhibit immune suppressive function. However, activin A positively regulates immune responses, as described in detail below. The activity
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of activin A in the immune cells partly overlaps with, but is clearly distinct from, that of TGF-b1. Immune cells may provide a model system for study the molecular basis of functional differences between activin and TGF-b.
II. Activin A in Th2 Cells A. Expression of activin A exclusively in T helper cells but not in regulatory T cells CD4þ T cells differentiate into T helper cells upon activation and play an important role in both humoral and cellular immune responses by producing multiple cytokines (Abbas et al., 1996). In murine CD4þ T cells, activin production is induced by their activation (Ogawa et al., 2006). Expression of the activin bA subunit, but not the bB subunit, has been detected in CD4þ T cells (Ogawa et al., 2006), indicating that they are able to produce activin A, but not activin AB or activin B. Increased production of activin A during CD4þ T cell activation is regulated at the mRNA level (Ogawa et al., 2006). CD4þ T cells have long been thought to differentiate only into effector T helper cells following their activation. However, over the past decade, a subpopulation of CD4þ T cells that constitutively expresses the IL-2 receptor a chain (CD25) has been identified as a unique subset of suppressor T cells. These CD4þ CD25þ regulatory T cells produce large amounts of TGF-b and are involved in maintaining peripheral immune tolerance (Asano et al., 1996; Sakaguchi, 2000; Shevach, 2002). They account for 7– 10% of all CD4þ T cells in normal mice (Papiernik et al., 1997). Interestingly, CD4þ CD25þ regulatory T cells from normal mice do not produce activin, even when they are fully activated by anti-CD3 and -CD28 crosslinking, whereas CD4þ CD25 conventional T helper cells produce high levels of activin following their activation (Ogawa et al., 2006). Also, while TGF-b1 inhibits T cell proliferation in a dose-dependent manner, activin A does not significantly affect T cell proliferation (Ogawa et al., 2006). These findings indicate that the expression and function of activin in CD4þ T cells are distinct from those of TGF-b.
B. Preferential production of activin A in Th2 cells In contrast to TGF-b1, which is produced in CD4þ CD25þ regulatory T cells (Asano et al., 1996; Sakaguchi, 2000; Shevach, 2002), activin A is exclusively produced in activated conventional CD4þ CD25 T helper cells (Ogawa et al., 2006). Furthermore, activin A is not involved in the suppressor function of T cells (Ogawa et al., 2006), suggesting that its roles in immune responses are distinct from those of TGF-b1. Upon antigen
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stimulation, naı¨ve CD4þ T cells differentiate into distinct populations of effector cells that exhibit characteristic patterns of cytokine production and immune regulation (Abbas et al., 1996). T helper 1 (Th1) cells secrete higher levels of interferon (IFN)-g and tumor necrosis factor (TNF)-a and are involved in cell-mediated immunity. In contrast, Th2 cells produce interleukin (IL)-4, IL-5, and IL-13 and are involved in humoral immune responses, promoting IgE production and eosinophil proliferation. In T helper cells, increased expression of activin A was found only in Th2 cells. When the differentiation of CD4þ T cells into T helper cell subsets was induced in vitro, higher activin secretion was detected in Th2 cells, but not Th1 cells (Ogawa et al., 2006). Expression of activin bA mRNA was high in Th2 cells, but not Th1 cells (Ogawa et al., 2006). Moreover, the secretion of activin was significantly higher in activated Th2 clones than Th1 clones. These results indicate that activin A production is associated with Th2-type immune responses. This conclusion is supported by a study showing that the secretion of activin A was increased in the airways of mice subjected to ovalbumin (OVA) sensitization followed by antigen challenge (Cho et al., 2003). Levels of activin A in bronchoalveolar lavage fluid from OVA-sensitized mice were also elevated after antigen challenge (Hardy et al., 2006). In a clinical study, serum levels of activin A were increased in patients with asthma (Karagiannidis et al., 2006). Moreover, T cells from these patients displayed elevated expression of activin bA mRNA.
C. Transactivation of the activin bA promoter by NFAT and c-Maf in Th2 cells Activin A production is markedly induced in cells that enter the Th2 pathway, suggesting that activin bA gene expression is regulated similarly to other Th2 cytokines. In fact, the mouse activin bA proximal promoter contains a binding site for c-Maf, a Th2-specific transcriptional factor (TGCTGATGTCA; nt 136 to 126) in close proximity to a nuclear factor of activated T cells (NFAT) binding site (TGGAAAAAC; nt 153 to 145) (Ogawa et al., 2006). c-Maf can synergize with NFAT to transactivate the activin bA gene, and, indeed, both factors have been implicated in activin bA gene transcription in Th2 cells (Ogawa et al., 2006). The cooperative regulation of activin bA gene expression by NFATp and c-Maf is consistent with the transcriptional regulation of the representative Th2 cytokine IL-4, in which c-Maf binds to a MARE immediately downstream of an NFAT site in the proximal IL-4 promoter and transactivates the IL-4 gene in synergy with NFATp (Ho et al., 1996). The findings of these studies indicate that activin A is a Th2 cytokine with a role in Th2 immune responses.
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III. Activin A in B Cells A. Production of activin A in activated B cells B cells are important effector cells in Th2 cell-mediated and humoral immune responses. Following their activation, resting B cells proliferate and differentiate into immunoglobulin-secreting plasma cells (DeFranco, 1987). In addition to antigens, various cytokines are involved in this process. Th1 cytokines are involved in cell-mediated immunity and immunoglobulin class switching to IgG2a. In contrast, Th2 cytokines are involved in humoral immune responses and immunoglobulin class switching, to IgG1 and IgE (Mosmann and Coffman, 1989; Mosmann and Sad, 1996). Activated B cells themselves also produce cytokines, some of which act as autocrine or paracrine factors. Activin A expression was induced in mouse splenic B cells in response to polyclonal activation with lipopolysaccharide (LPS) (Ogawa et al., 2008). Expression of follistatin has also been detected in B cells (Ogawa et al., 2008). Follistatin is an activin-binding protein that neutralizes activin activity in many biological systems (Nakamura et al., 1990; Woodruff, 1998). Interestingly, the expression of follistatin is inversely related to that of activin bA in B cell activation: B cell expression of activin bA and follistatin are increased and decreased, respectively, in response to activation (Ogawa et al., 2008). The increased net activin activity in the culture supernatant of LPS-stimulated B cells can be explained by a combination of increased activin A production and decreased follistatin production. The findings of this last study suggest the involvement of follistatin in the regulation of activin function in the immune system.
B. Involvement of activin A in immunoglobulin production in B cells Activin has been shown to induce growth arrest and apoptosis in a B cellderived cell line (Yamato et al., 1997) and B cell hybridomas (Hashimoto et al., 1998). Moreover, although activin A inhibited the generation of B cells from marrow stem cells (Zipori and Barda-Saad, 2001), it stimulated proliferation and antibody secretion in mature B cells (Ogawa et al., 2008), suggesting that the effects of activin A on B cells are stage-dependent. A study performed using human peripheral blood mononuclear cells (PBMCs) showed that activin A stimulated IgE production in PBMCs cultured in the presence of IL-4 (Yamashita et al., 1993). However, it did not influence IgE production in purified B cells, even in the presence of an anti-CD40 mAb and IL-4 (Yamashita et al., 1993). In an in vivo study, the neutralization of circulating activin A decreased the strength of an antigenspecific IgE response (Ogawa et al., 2008). Activin A, produced by activated
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B cells, is involved in antigen-specific IgE production, presumably through the activation of other immune cells. Separately, neutralization of activin A resulted in a significant decrease in serum IL-4 levels in OVA-immunized mice (Ogawa et al., 2008), suggesting that this response may be realized through the regulation of IL-4 production. In addition to activated B cells, activin A is produced by other immune cells involved in Th2 immune responses, including Th2 cells (Ogawa et al., 2006), macrophages (Ogawa et al., 2000), dendritic cells (Robson et al., 2008), and mast cells (Funaba et al., 2003a). These cells may also regulate B cell function by secreting activin A. Thus, activin A stimulates Th2-mediated immune responses by enhancing antibody production. TGF-b, the prototype of the TGF-b superfamily of proteins, inhibits proliferation and induces apoptosis in immature and resting B cells (Li et al., 2006). In contrast, it also positively regulates B cell responses by inducing class switching to IgA and IgG2b (Li et al., 2006). In most in vitro studies, the biological activities of activin overlap with those of TGF-b (Lebrun et al., 1997). This is partly due to the fact that activin and TGF-b activate the same signaling intermediates: Smad2 and/or Smad3 (Massague´ and Chen, 2000). It would, thus, not be unexpected if activin A had similar effects to TGF-b on B cells. However, activin A does not induce immunoglobulin class switching to IgA and IgG2b in B cells (Ogawa et al., 2008). Thus, the function of activin A in B cells is quite different to that of TGF-b, suggesting that these two structurally related proteins perform different roles in the immune system. So far, there are no known differences between the signaling pathways activated by activin and TGF-b. B cells may provide a good model system for studying the molecular bases of the functional differences between activin and TGF-b.
IV. Activin A in Macrophages A. Activation-induced expression of activin A in macrophages Macrophages are critical effector cells involved in both innate and adaptive immune responses. Activated macrophages produce many cytokines and chemokines that play pivotal roles in inflammatory processes (Mosser, 2003). Various studies have shown that monocyte-/macrophage-lineage cells produce activin A. Human peripheral blood monocytes produce activin A, and its expression is increased following their activation (Eramaa et al., 1992; Shao et al., 1992; Yu et al., 1996). Mouse alveolar macrophages also express activin A (Matsuse et al., 1995). Activin production and activin bA mRNA levels in mouse peritoneal macrophages were increased following their activation by LPS (Ogawa et al., 2000). Human monocyte-derived dendritic cells and peripheral blood myeloid dendritic
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cells too express activin A and rapidly secrete it in large amounts following exposure to bacteria, specific Toll-like receptor ligands, or CD40 ligand (Robson et al., 2008). Changes in the expression of activin in the monocyte/macrophage lineage are different from those of TGF-b gene transcripts. While TGF-b 1 mRNA expression in human monocytes was found not to be affected by LPS stimulation in one study (Assoian et al., 1987), a second study reported TGF-b1 mRNA levels to decrease rapidly (within 3 h) following the treatment of peripheral blood mononuclear cells with LPS (Zhou et al., 1992). Furthermore, LPS treatment strongly suppressed TGF-b1 mRNA expression in peritoneal macrophages (Ogawa et al., 2011). Their distinct patterns of expression in response to LPS stimulation suggest that activin and TGF-b perform different functions in macrophages.
B. Production of type IV collagenase in macrophages stimulated by activin A Macrophages regulate the inflammatory response in innate immunity through their production of matrix metalloproteinases (MMPs), a major family of matrix-degrading enzymes (Nagase and Woessner, 1999; Owen and Campbell, 1999). MMPs share certain biochemical properties, yet each has a distinct substrate specificity. The type IV collagenases, MMP2 (72 kDa gelatinase A) and MMP-9 (92 kDa gelatinase B), are important members of the MMP family and are involved in the cleavage of denatured collagens of all types and native basement membrane components (Nagase and Woessner, 1999; Owen and Campbell, 1999). Expression of type IV collagenases is regulated by a variety of factors, including cytokines, growth factors, chemical agents, physical stress, and cell transformation (Mauviel, 1993). LPS stimulates the production of MMP-2 and MMP-9 in macrophages (Owen and Campbell, 1999). Activin has been shown to induce the expression and activation of MMP-2 in mouse peritoneal macrophages, increasing MMP-2 mRNA level net gelatinolytic activity without affecting MMP-9 expression (Ogawa et al., 2000). LPS-stimulated MMP-2 production, while exogenous activin A further increased MMP-2 expression in LPS-stimulated peritoneal macrophages (Ogawa et al., 2000). Similar observations have been made in other cells. For example, activin A specifically induced the early expression of MMP-2 within villous cytotrophoblast cells (Caniggia et al., 1997) and stimulated endometrial production of pro-MMP-2 and active MMP-2 ( Jones et al., 2006). These effects on macrophages partly overlap with, but are clearly distinct from, those of TGF-b1. TGF-b was shown to stimulate the production of both MMP-2 and MMP-9 in mouse peritoneal macrophages, but blocked LPS-induced MMP-9 expression (Xie et al., 1994). Considering that MMP-2 is the rate-limiting enzyme in the degradation of basement membrane
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collagens (Welgus et al., 1990), activin A may influence the migration and infiltration of macrophages through the basement membrane in inflammatory states. This would represent a different role from that of TGF-b, which is well known to be an anti-inflammatory cytokine (Bogdan et al., 1992; Shull et al., 1992; Tsunawaki et al., 1988; Williams et al., 1996; Yaswen et al., 1996).
C. Alternative activation of macrophages induced by activin A Macrophages play important roles in Th1 and Th2 immune responses (Mirmonsef et al., 1999). In parallel with the separation of T helper cells into Th1 and Th2 cells, the exposure of macrophages to a specific set of cytokines likewise biases them toward either an M1 (classically activated) or M2 (alternatively activated) phenotype (Dickensheets and Donnelly, 1999; Goerdt et al., 1999; Mills et al., 2000). Classical activation of macrophages with Th1 cytokines results in free radical release and increased cytokine secretion, essential components of a successful response to infection by intracellular bacteria and viruses (Heinzel et al., 1989; Scott et al., 1988). In contrast, the alternative activation of macrophages with Th2 cytokines is required for defense against extracellular pathogens and parasites (Finkelman and Urban, 2001). Activated macrophages metabolize L-arginine by two alternative pathways involving the enzymes iNOS and arginase (Granger et al., 1990; Modolell et al., 1995). Th1 cytokines induce macrophages to produce iNOS, whereas Th2 cytokines induce arginase-1 production (Corraliza et al., 1995; Modolell et al., 1995; Munder et al., 1998). While iNOS converts L-arginine into NO and L-citrulline, arginase catalyzes its metabolism to urea and L-ornithine. Thus, iNOS and arginase-1 effectively compete for L-arginine, and thereby negatively regulate each other’s function. Like other Th2 cytokines, activin A inhibited IFN-g-induced NO2 production in both peritoneal and RAW264.7 macrophages (Ogawa et al., 2006). The mRNA expression of arginase-1, a marker of alternatively activated macrophages (M2 phenotype), was clearly augmented in macrophages following treatment with activin A (Ogawa et al., 2006). In contrast, IFN-ginduced expression of iNOS, a marker of classically activated macrophages (M1 phenotype), was decreased in macrophages treated with activin A (Ogawa et al., 2006). These observations functionally implicate activin A in the generation of alternatively activated macrophages (Fig. 12.1).
V. Activin A in Mast Cells A. Upregulation of activin A expression in activated mast cells The multivalent binding of antigen to receptor-bound IgE and the subsequent aggregation of the high-affinity IgE receptor (FceRI) provide the trigger for the activation of mast cells, leading to an immune response
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Naive macrophages Activin A
Activin A
Inhibits
Promotes
M1 phenotype Classically activated macrophages
M2 phenotype Alternatively activated macrophages
iNOS
Arginase-1 l-arginine
NO and l-citrulline Th1-type immune responses Intracellular pathogens
Urea and l-ornithine Th2-type immune responses Extracellular pathogens
Figure 12.1 Schematic representation of the proposed roles of activin A in the activation of macrophages. Similar to other Th2 cytokines, such as IL-4 and IL-13, activin A promotes the alternative activation of macrophages, while inhibiting their classical activation.
(Kalesnikoff and Galli, 2008; Rivera and Olivera, 2007). Mast cells are produced in the bone marrow as progenitor cells. Committed mast cell precursors then migrate into the peripheral blood, from where they invade target tissues (Galli et al., 2008a; Kitamura et al., 1993). Mast cells mature in target tissues under the regulation of local growth and differentiation factors, acquiring a characteristic morphology characterized by metachromatic granules, and displaying increased expression of mast cell proteases (MCPs) (Galli et al., 2008a; Gurish and Boyce, 2002). Activin A expression has been shown to increase in activated mast cells. Upregulation of activin bA, but not bB, mRNA, and protein expression in mast cells was observed in response to IgE- and antigen-dependent stimulation (Cho et al., 2003; Funaba et al., 2003a). The engagement of antigen receptors initiates a cascade of protein tyrosine kinase phosphorylation events, leading to an increase in cytosolic Ca2þ levels (due to release from intracellular stores) (Galli et al., 2008b; Kalesnikoff and Galli, 2008; Rivera and Olivera, 2007). The emptying of intracellular Ca2þ stores drives the influx of extracellular Ca2þ in a process known as store-operated Ca2þ influx (Kalesnikoff and Galli, 2008; Rivera and Olivera, 2007). The increase in activin bA mRNA expression was reproduced by treatment with ionomycin, a Ca2þ ionophore, in a response that required de novo protein synthesis (Cho et al., 2003; Funaba et al., 2003a). The Ca2þ-induced activation of calmodulin-dependent kinase and calcineurin was found to be responsible for the upregulation of activin bA expression, which also involved the activation of JNK and p38 MAP kinase (Funaba et al., 2003a). In view of the findings in Th2 cells (Ogawa et al., 2006), B cells (Ogawa et al., 2008), macrophages (Ogawa et al., 2000), and dendritic cells (Robson
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et al., 2008) described above, the upregulation of activin A expression in response to cellular activation may be a general feature of immune cells.
B. Modulation of mast cell function by activin A Mast cells are one of the target cells for activin A secreted from activated immune cells. Activin A has been shown to modulate the morphology and function of bone marrow-derived mast cells (BMMCs) in vitro. Treatment with activin A increased the number of BMMCs containing strongly metachromatic granules (Funaba et al., 2003b), and showing a round, compact morphology. In fact, the reduction in nuclear size was greater than that of the cytoplasm, resulting in a decrease in the nuclear– cytoplasmic ratio. These features recall the morphometric characteristics of mature mast cells (Craig et al., 1989), suggesting a stimulatory role for activin A in mast cell maturation. The maturation of mast cells is also characterized by the synthesis of MCPs (Kalesnikoff and Galli, 2008; Miller and Pemberton, 2002). Treatment with activin A increases BMMC expression of several MCPs, including MCP-1 (Funaba et al., 2003b), MCP-6 (Funaba et al., 2005), and MCP-7 (Funaba et al., 2003c). Of these three proteases, activin A regulated the expression of two, MCP-6 and MCP-7, at the level of transcription, responses for which Smad3, but not Smad2, was responsible (Funaba et al., 2003c, 2005). Transcription of the genes encoding MCP-6 and MCP-7 was also modulated by microphthalmia-associated transcription factor (MITF), which is predominantly expressed in mast cells (Kitamura et al., 2000). The Mad homology 1 (MH1) domain and linker region of Smad3 were shown to associate with the carboxyl terminus of MITF (Funaba et al., 2003c). This association differentially affected the transcription of the genes encoding MCP-6 and MCP-7. While MITF enhanced Smad3-mediated transcription of MCP-6 (Funaba et al., 2005), it blocked Smad3-mediated transcription of MCP-7 (Funaba et al., 2003c). This latter inhibitory effect was partly realized through the degradation of Smad3 protein. Reports describing the effects of activin A on mast cell growth are conflicting. The addition of activin A to the culture medium dose-dependently inhibited the proliferation of BMMC, suggesting that it inhibited mast cell growth (Funaba et al., 2003b). This response may relate to the promotion of mast cell maturation by activin A. However, the results of a second study suggest that activin A may stimulate mast cell proliferation. The doubling time of BMMCs prepared from Smad3-null mice was longer than that of cells from wild-type mice, as a result of increased cell death (Funaba et al., 2006a). Basal activin activity may thus be necessary for BMMC survival.
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In addition to its regulatory effects on mast cell maturation, activin A also acts as a chemoattractant for BMMCs. Activin A was shown to chemoattract cultured BMMCs, dose-dependently (Funaba et al., 2003b). Concentrations of activin A required to induce migration (10 pM) were lower than those needed to upregulate MCP expression ( 1 nM) or inhibit cell growth (IC50 ¼ 2 nM). The graph of BMMC migration against activin A concentration has a bell-shaped curve. Thus, an activin A concentration gradient may initially serve to chemoattract circulating mast cell precursors. As these precursors gradually move closer to the activin A source, and the local activin A concentration consequently increases, they may subsequently mature (Fig. 12.2). Unlike in Th2 cells (Ogawa et al., 2006), B cells (Ogawa et al., 2008), and peritoneal macrophages (Ogawa et al., 2000, 2011), the roles of activin A in mast cells are similar to those of TGF-b. Morphological changes, upregulation of MCP expression, cell growth inhibition, and the
The site of inflammation Activin A Activin A concentration gradient proximal Activated mast cells High conc (~1 nM) Matured mast cells
Maturation
Low conc (~10 pM) Mast cell precursors
Chemotactic migration
Distal
Peripheral blood circulation
Figure 12.2 Schematic representation of the proposed roles of activin A in mast cell function. Activin A produced by activated immune cells at sites of inflammation may form a concentration gradient through diffusion. Activin A at concentrations as low as 10 pM may act as chemoattractant of circulating mast cell precursors. The mast cell precursors subsequently mature as the local activin A concentration increases (to approximately 1 nM).
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stimulation of directional migration are all responses that are shared with TGF-b (Funaba et al., 2003b,c, 2005, 2006b; Miller et al., 1999; Olsson et al., 2000, 2001). The diverse effects of TGF-b superfamily members are realized through the activation of Smad-dependent and -independent pathways (Feng and Derynck, 2005). As mentioned above, activin A- and TGFb-induced Smad signaling pathways are indistinguishable from one another (Feng and Derynck, 2005). Thus, the functional effects of activin A on mast cells may be largely mediated by a Smad-dependent pathway. Smad3-null BMMCs were unable to migrate to TGF-b (Funaba et al., 2006b). In addition to their IgE-dependent responses, mast cells function as effector cells during innate immune responses (Dawicki and Marshall, 2007). Bacterial infection stimulates the synthesis and secretion of a wide spectrum of mediators and cytokines in mast cells (Dawicki and Marshall, 2007). BMMCs from Smad3-null mice displayed enhanced production of proinflammatory cytokines in response to LPS (Kanamaru et al., 2005). In a mouse model of acute septic peritonitis (induced through cecal ligation and puncture), the survival rate for mast cell-deficient W/WV mice replenished with Smad3-deficient BMMCs was higher than that for animals replenished with wild-type BMMCs (Kanamaru et al., 2005). These findings suggest that Smad3 negatively regulated mast cell-mediated innate immune responses to Gram-negative bacteria.
VI. Conclusions and Future Directions The expression of activin A, but not activin AB or activin B, has been detected in many types of immune cells, including Th2 cells (Ogawa et al., 2006), B cells (Ogawa et al., 2008), alveolar macrophages (Matsuse et al., 1995), peritoneal macrophages (Ogawa et al., 2000), monocyte-derived dendritic cells (Robson et al., 2008), peripheral blood myeloid dendritic cells (Robson et al., 2008), and mast cells (Funaba et al., 2003a). Thus, activin A may be the major activin in immune cells. Indeed, upregulation of activin A expression appears to be a general feature of immune cell activation. Although many studies have demonstrated the induction of activin A expression, the in vivo function of activin A in the immune system has not yet been fully elucidated. This is partly due to the lack of a suitable in vivo model. Activin bA-deficient mice die within 24 h of birth (Matzuk et al., 1995a,b). The results of many studies indicate that activin A positively regulates Th2 immune responses. However, conflicting results have also been reported. Activin A was able to promote the conversion of conventional CD4þ CD25 T helper cells into regulatory T cells in vitro (Huber et al., 2009). Furthermore, it suppressed Th2 responses by inducing the
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production of antigen-specific regulatory T cells (Semitekolou et al., 2009). Thus, the potential role of activin A in the immune system in vivo remains controversial and must be examined in future studies. Each member of the TGF-b superfamily initiates responses by binding to a characteristic combination of type I and II receptors, both of which are needed for signaling (Wrana et al., 1994). After ligand binding, the activated type I receptor relays the signal to receptor-regulated Smads (R-Smads) by phosphorylating them at sites in their C-termini (Abdollah et al., 1997; Souchelnytskyi et al., 1997). Upon phosphorylation, the R-Smads form hetero-oligomeric complexes with Smad4, the common mediator Smad (Hahn et al., 1996; Lagna et al., 1996). These complexes then migrate to the nucleus and activate gene transcription, through direct DNA binding or by associating with other sequence-specific transcription factors (Massague´ and Wotton, 2000). Activin and TGF-b signal through the same R-Smads, Smad2 and/or Smad3 (Lebrun et al., 1997; Massague´ and Chen, 2000), and consequently have overlapping biological activities in many different types of cells. However, as shown in Table 12.1, murine activin A and TGF-b1 share only 27.2% amino acid sequence homology. The sequence homology between their respective type II receptors is also relatively low (<32%). Thus, they are structurally distinct from each other. How these two distinct proteins have evolved to serve the same function remains unclear. As mentioned above, the expression pattern of activin A in immune cells is different from that of TGF-b1, indicating that they perform different functions in the immune system. In fact, the activity of activin A in immune cells partly overlaps with, but is clearly distinct from, that of TGF-b1. This discrepancy suggests that a signaling pathway not involving Smad2 or Smad3 is likely responsible for the differences between activin A and TGF-b1 activities in immune cells. Immune cells may provide a model system for studying the molecular bases of functional differences between activin A and TGF-b1. Table 12.1
Sequence homology between of TGF-b and activin signaling molecules
TGF-b signal
Ligands Type II Rs Type I Rs R-Smads
TGF-b1 (390) TbRII (592)
Activin signal
Activin A (424) ActRII (513) ActRIIB (536) ALK5 (503) ALK4 (505) Smad2 (467) and Smad3 (425)
Amino acid identity (%)
27.2 31.9 31.6 69.6 100
Type II Rs, type II receptors; type I Rs, type I receptors; R-Smads, receptor-regulated Smads numbers indicate protein sizes (number of amino acids).
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REFERENCES Abbas, A. K., Murphy, K. M., and Sher, A. (1996). Functional diversity of helper T lymphocytes. Nature 383, 787–793. Abdollah, S., Macias-Silva, M., Tsukazaki, T., Hayashi, H., Attisano, L., and Wrana, J. L. (1997). TbRI phosphorylation of Smad2 on Ser465 and Ser467 is required for Smad2– Smad4 complex formation and signaling. J. Biol. Chem. 272, 27678–27685. Asano, M., Toda, M., Sakaguchi, N., and Sakaguchi, S. (1996). Autoimmune disease as a consequence of developmental abnormality of a T cell subpopulation. J. Exp. Med. 184, 387–396. Assoian, R. K., Fleurdelys, B. E., Stevenson, H. C., Miller, P. J., Madtes, D. K., Raines, E. W., Ross, R., and Sporn, M. B. (1987). Expression and secretion of type b transforming growth factor by activated human macrophages. Proc. Natl. Acad. Sci. USA 84, 6020–6024. Bilezikjian, L. M., Corrigan, A. Z., and Vale, W. (1990). Activin A modulates growth hormone secretion from cultures of rat anterior pituitary cells. Endocrinology 126, 2369. Bogdan, C., Paik, J., Vodovotz, Y., and Nathan, C. (1992). Contrasting mechanisms for suppression of macrophages cytokine release by transforming growth factor-b and interleukin-10. J. Biol. Chem. 267, 23301–23308. Caniggia, I., Lye, S. J., and Cross, J. C. (1997). Activin is a local regulator of human cytotrophoblast cell differentiation. Endocrinology 138, 3976–3986. Chen, Y., Kam, C. S., Liu, F. Q., Liu, Y., Lui, V. C., Lamb, J. R., and Tam, P. K. (2008). LPS-induced up-regulation of TGF-b receptor 1 is associated with TNF-a expression in human monocyte-derived macrophages. J. Leukoc. Biol. 83, 1165–1173. Cho, S. H., Yao, Z., Wang, S. W., Alban, R. F., Barbers, R. G., French, S. W., and Oh, C. K. (2003). Regulation of activin A expression in mast cells and asthma: Its effect on the proliferation of human airway smooth muscle cells. J. Immunol. 170, 4045–4052. Corraliza, I. M., Soler, G., Eichmann, K., and Modolell, M. (1995). Arginase induction by suppressors of nitric oxide synthesis (IL-4, IL-10 and PGE2) in murine bone marrowderived macrophages. Biochem. Biophys. Res. Commun. 206, 667–673. Craig, S. S., Schechter, N. M., and Schwartz, L. B. (1989). Ultrastructural analysis of maturing human T and TC mast cells in situ. Lab. Invest. 60, 147–157. Dawicki, W., and Marshall, J. S. (2007). New and emerging roles for mast cells in host defence. Curr. Opin. Immunol. 19, 31–38. DeFranco, A. L. (1987). Molecular aspects of B-lymphocyte activation. Annu. Rev. Cell Biol. 3, 143–178. Dennler, S., Goumans, M. J., and ten Dijke, P. (2002). Transforming growth factor b signal transduction. J. Leukoc. Biol. 71, 731–740. Dickensheets, H. L., and Donnelly, R. P. (1999). Inhibition of IL-4-inducible gene expression in human monocytes by type I and type II interferons. J. Leukoc. Biol. 65, 307–312. Eramaa, M., Hurme, M., Stenman, U. H., and Ritvos, O. (1992). Activin A/erythroid differentiation factor is induced during human monocyte activation. J. Exp. Med. 176, 1449–1452. Feng, X. H., and Derynck, R. (2005). Specificity and versatility in TGF-b signaling through Smads. Annu. Rev. Cell Dev. Biol. 21, 659–693. Finkelman, F. D., and Urban, J. F. (2001). The other side of the coin: The protective role of the TH2 cytokines. J. Allergy Clin. Immunol. 107, 772–780. Funaba, M., Ikeda, T., Ogawa, K., and Abe, M. (2003a). Calcium-regulated expression of activin A in RBL-2H3 mast cells. Cell. Signal. 15, 605–613. Funaba, M., Ikeda, T., Ogawa, K., Murakami, M., and Abe, M. (2003b). Role of activin A in murine mast cells: Modulation of cell growth, differentiation, and migration. J. Leukoc. Biol. 73, 793–801.
Activin in Humoral Immune Responses
249
Funaba, M., Ikeda, T., Murakami, M., Ogawa, K., Tsuchida, K., Sugino, H., and Abe, M. (2003c). Transcriptional activation of mouse mast cell protease-7 by activin and transforming growth factor-b is inhibited by microphthalmia-associated transcription factor. J. Biol. Chem. 278, 52032–52041. Funaba, M., Ikeda, T., Murakami, M., Ogawa, K., and Abe, M. (2005). Up-regulation of mouse mast cell protease-6 gene by transforming growth factor-b and activin in mast cell progenitors. Cell. Signal. 17, 121–128. Funaba, M., Nakaya, K., Ikeda, T., Murakami, M., Tsuchida, K., and Sugino, H. (2006a). Requirement of Smad3 for mast cell growth. Cell. Immunol. 240, 47–52. Funaba, M., Ikeda, T., Murakami, M., Ogawa, K., Nishino, Y., Tsuchida, K., Sugino, H., and Abe, M. (2006b). Involvement of p38 MAP kinase and Smad3 in TGF-b-mediated mast cell functions. Cell. Signal. 18, 2154–2161. Galli, S. J., Grimbaldeston, M., and Tsai, M. (2008a). Immunomodulatory mast cells: Negative, as well as positive, regulators of immunity. Nat. Rev. Immunol. 8, 478–486. Galli, S. J., Tsai, M., and Piliponsky, A. M. (2008b). The development of allergic inflammation. Nature 454, 445–454. Goerdt, S., Politz, O., Schledzewski, K., Birk, R., Gratchev, A., Guillot, P., Hakiy, N., Klemke, C. D., Dippel, E., Kodelja, V., and Orfanos, C. E. (1999). Alternative versus classical activation of macrophages. Pathobiology 67, 222–226. Granger, D. L., Hibbs, J. B., Perfect, J. R., and Durack, D. T. (1990). Metabolic fate of L-arginine in relation to microbiostatic capability of murine macrophages. J. Clin. Invest. 85, 264–273. Gurish, M. F., and Boyce, J. A. (2002). Mast cell growth, differentiation, and death. Clin. Rev. Allergy Immunol. 22, 107–118. Hahn, S. A., Schutte, M., Hoque, A. T., Moskaluk, C. A., da Costa, L. T., Rozenblum, E., Weinstein, C. L., Fischer, A., Yeo, C. J., Hruban, R. H., and Kern, S. E. (1996). DPC4, a candidate tumor suppressor gene at human chromosome 18q21.1. Science 271, 350–353. Hardy, C. L., O0 Connor, A. E., Yao, J., Sebire, K., de Kretser, D. M., Rolland, J. M., Anderson, G. P., Phillips, D. J., and O0 Hehir, R. E. (2006). Follistatin is a candidate endogenous negative regulator of activin A in experimental allergic asthma. Clin. Exp. Allergy 36, 941–950. Hashimoto, M., Kondo, S., Sakurai, T., Etoh, Y., Shibai, H., and Muramatsu, M. (1990). Activin/EDF as an inhibitor of neural differentiation. Biochem. Biophys. Res. Commun. 173, 193. Hashimoto, O., Yamato, K., Koseki, T., Ohguchi, M., Ishisaki, A., Shoji, H., Nakamura, T., Hayashi, Y., Sugino, H., and Nishihara, T. (1998). The role of activin type I receptors in activin A-induced growth arrest and apoptosis in mouse B-cell hybridoma cells. Cell. Signal. 10, 743–749. Heinzel, F. P., Sadick, M. D., Holaday, B. J., Coffman, R. L., and Locksley, R. M. (1989). Reciprocal expression of interferon gamma or interleukin 4 during the resolution or progression of murine leishmaniasis. Evidence for expansion of distinct helper T cell subsets. J. Exp. Med. 169, 59–72. Hemmati-Brivanlou, A., and Melton, D. A. (1994). Inhibition of activin receptor signaling promotes neuralization in Xenopus. Cell 77, 273. Hemmati-Brivanlou, A., Kelly, O. G., and Melton, D. A. (1994). Follistatin, an antagonist of activin, is expressed in the Spemann organizer and displays direct neuralizing activity. Cell 77, 283. Ho, I. C., Hodge, M. R., Rooney, J. W., and Glimcher, L. H. (1996). The proto-oncogene c-maf is responsible for tissue-specific expression of interleukin-4. Cell 85, 973–983. Huber, S., Stahl, F. R., Schrader, J., Lu¨th, S., Presser, K., Carambia, A., Flavell, R. A., Werner, S., Blessing, M., Herkel, J., and Schramm, C. (2009). Activin a promotes the
250
Kenji Ogawa and Masayuki Funaba
TGF-b-induced conversion of CD4þCD25 T cells into Foxp3þ induced regulatory T cells. J. Immunol. 182, 4633–4640. Jones, R. L., Findlay, J. K., Farnworth, P. G., Robertson, D. M., Wallace, E., and Salamonsen, L. A. (2006). Activin A and inhibin A differentially regulate human uterine matrix metalloproteinases: Potential interactions during decidualization and trophoblast invasion. Endocrinology 147, 724–732. Kalesnikoff, J., and Galli, S. J. (2008). New developments in mast cell biology. Nat. Immunol. 9, 1215–1223. Kanamaru, Y., Sumiyoshi, K., Ushio, H., Ogawa, H., Okumura, K., and Nakao, A. (2005). Smad3 deficiency in mast cells provides efficient host protection against acute septic peritonitis. J. Immunol. 174, 4193–4197. Karagiannidis, C., Hense, G., Martin, C., Epstein, M., Ruckert, B., Mantel, P. Y., Menz, G., Uhlig, S., Blaser, K., and Schmidt-Weber, C. B. (2006). Activin A is an acute allergen-responsive cytokine and provides a link to TGF-b-mediated airway remodeling in asthma. J. Allergy Clin. Immunol. 117, 111–118. Kitamura, Y., Kasugai, T., Arizono, N., and Matsuda, H. (1993). Development of mast cells and basophils: Processes and regulation mechanisms. Am. J. Med. Sci. 306, 185–191. Kitamura, Y., Morii, E., Jippo, T., and Ito, A. (2000). Mi-transcription factor as a regulator of mast cell differentiation. Int. J. Hematol. 71, 197–202. Lagna, G., Hata, A., Hemmati-Brivanlou, A., and Massague´, J. (1996). Partnership between DPC4 and SMAD proteins in TGF-b signalling pathways. Nature 383, 832–836. Lebrun, J. J., Chen, Y., and Vale, W. (1997). Receptor serine kinases and signaling by activins and inhibins. In “Inhibin, Activin and Follistatin: Regulatory Functions in System and Cell Biology,” (T. Aono, H. Sugino, and W. W. Vale, Eds.), pp. 1–20. Springer-Verlag, New York. Letterio, J. J. (2000). Murine models define the role of TGF-b as a master regulator of immune cell function. Cytokine Growth Factor Rev. 11, 81–87. Li, W., Yuen, B. H., and Leung, P. C. (1992). Inhibition of progestin accumulation by activin-A in human granulosa cells. J. Clin. Endocrinol. Metab. 75, 285–289. Li, M. O., Wan, Y. Y., Sanjabi, S., Robertson, A. K., and Flavell, R. A. (2006). Transforming growth factor-b regulation of immune responses. Annu. Rev. Immunol. 24, 99–146. Massague´, J., and Chen, Y. G. (2000). Controlling TGFb signaling. Genes Dev. 14, 627–644. Massague´, J., and Wotton, D. (2000). Transcriptional control by the TGF-b/Smad signaling system. EMBO J. 19, 1745–1754. Matsuse, T., Fukuchi, Y., Eto, Y., Matsui, H., Hosoi, T., Oka, T., Ohga, E., Nagase, T., and Orimo, H. (1995). Expression of immunoreactive and bioactive activin A protein in adult murine lung after bleomycin treatment. Am. J. Respir. Cell Mol. Biol. 13, 17–24. Matzuk, M. M., Kumar, T. R., and Bradley, A. (1995a). Different phenotypes for mice deficient in either activins or activin receptor type II. Nature 374, 356–360. Matzuk, M. M., Kumar, T. R., Vassalli, A., Bickenbach, J. R., Roop, D. R., Jaenisch, R., and Bradley, A. (1995b). Functional analysis of activins during mammalian development. Nature 374, 354–356. Mauduit, C., Chauvin, M. A., dePeretti, E., Morera, A. M., and Benahmed, M. (1991). Effect of activin A on dehydroepiandrosterone and testosterone secretion by primary immature porcine Leydig cells. Biol. Reprod. 45, 101–109. Mauviel, A. (1993). Cytokine regulation of metalloproteinase gene expression. J. Cell. Biochem. 53, 288–295. Miller, H. R., and Pemberton, A. D. (2002). Tissue-specific expression of mast cell granule serine proteinases and their role in inflammation in the lung and gut. Immunology 105, 375–390. Miller, H. R., Wright, S. H., Knight, P. A., and Thornton, E. M. (1999). A novel function for transforming growth factor-b1: Upregulation of the expression and the IgE-
Activin in Humoral Immune Responses
251
independent extracellular release of a mucosal mast cell granule-specific b-chymase, mouse mast cell protease-1. Blood 93, 3473–3486. Mills, C. D., Kincaid, K., Alt, J. M., Heilman, M. J., and Hill, A. M. (2000). M-1/M-2 macrophages and the Th1/Th2 paradigm. J. Immunol. 164, 6166–6173. Mirmonsef, P., Shelburne, C. P., Fitzhugh, Y. C., Chong, H. J., and Ryan, J. J. (1999). Inhibition of Kit expression by IL-4 and IL-10 in murine mast cells: Role of STAT6 and phosphatidylinositol 30 -kinase. J. Immunol. 163, 2530–2539. Miro, F., Smyth, C. D., and Hillier, S. G. (1991). Development-related effects of recombinant activin on steroid synthesis in rat granulosa cells. Endocrinology 129, 3388–3394. Modolell, M., Corraliza, I. M., Link, F., Soler, G., and Eichmann, K. (1995). Reciprocal regulation of nitric oxide synthase/arginase balance in mouse bone marrow-derived macrophages by Th1 and Th2 cytokines. Eur. J. Immunol. 25, 1101–1104. Mosmann, T. R., and Coffman, R. L. (1989). TH1 and TH2 cells: Different patterns of lymphokine secretion lead to different functional properties. Annu. Rev. Immunol. 7, 145–173. Mosmann, T. R., and Sad, S. (1996). The expanding universe of T-cell subsets: Th1 Th2 and more. Immunol. Today 17, 138–146. Mosser, D. M. (2003). The many faces of macrophage activation. J. Leukoc. Biol. 73, 209–212. Munder, M., Eichmann, K., and Modolell, M. (1998). Alternative metabolic states in murine macrophages reflected by the nitric oxide synthase/arginase balance: Competitive regulation by CD4þ T cells correlates with Th1/Th2 phenotype. J. Immunol. 160, 5347–5354. Murata, T., and Ying, S.-Y. (1991). Transforming growth factor-b and activin inhibit basal secretion of prolactin in pituitary monolyer culture system. Proc. Soc. Exp. Biol. Med. 198, 599–605. Nagase, H., and Woessner, J. F., Jr. (1999). Matrix metalloproteinases. J. Biol. Chem. 274, 21491–21494. Nakamura, T., Takio, K., Eto, Y., Shibai, H., Titani, K., and Sugino, H. (1990). Activinbinding protein from rat ovary is follistatin. Science 247, 836–838. Ogawa, K., Funaba, M., Mathews, L. S., and Mizutani, T. (2000). Activin A stimulates type IV collagenase (matrix metalloproteinase-2) production in mouse peritoneal macrophages. J. Immunol. 165, 2997–3003. Ogawa, K., Funaba, M., Chen, Y., and Tsujimoto, M. (2006). Activin A functions as a Th2 cytokine in the promotion of the alternative activation of macrophages. J. Immunol. 177, 6787–6794. Ogawa, K., Funaba, M., and Tsujimoto, M. (2008). A dual role of activin A in regulating immunoglobulin production of B cells. J. Leukoc. Biol. 83, 1451–1458. Ogawa, K., Funaba, M., and Tsujimoto, M. (2011). The effect of TGF-b1 on the expression of type IV collagenases in mouse peritoneal macrophages. Mol. Biol. Rep. (in press). Olsson, N., Piek, E., ten Dijke, P., and Nilsson, G. (2000). Human mast cell migration in response to members of the transforming growth factor-b family. J. Leukoc. Biol. 67, 350–356. Olsson, N., Piek, E., Sundstro¨m, M., ten Dijke, P., and Nilsson, G. (2001). Transforming growth factor-b-mediated mast cell migration depends on mitogen-activated protein kinase activity. Cell. Signal. 13, 483–490. Owen, C. A., and Campbell, E. J. (1999). The cell biology of leukocyte-mediated proteolysis. J. Leukoc. Biol. 65, 137–150. Papiernik, M., Leite de Moraes, M. C., Pontoux, C., Joret, A.-M., Rocha, B., Penit, C., and Dy, M. (1997). T cell deletion induced y chronic infection with mouse mammary tumor virus spares a CD25-positive, IL-10-producing T cell population with infectious capacity. J. Immunol. 158, 4642–4653.
252
Kenji Ogawa and Masayuki Funaba
Rivera, J., and Olivera, A. (2007). Src family kinases and lipid mediators in control of allergic inflammation. Immunol. Rev. 217, 255–268. Robson, N. C., Phillips, D. J., McAlpine, T., Shin, A., Svobodova, S., Toy, T., Pillay, V., Kirkpatrick, N., Zanker, D., Wilson, K., Helling, I., Wei, H., et al. (2008). Activin-A: A novel dendritic cell-derived cytokine that potently attenuates CD40 ligand-specific cytokine and chemokine production. Blood 111, 2733–2743. Sakaguchi, S. (2000). Regulatory T cells: Key controllers of immunologic self-tolerance. Cell 101, 455–458. Schubert, D., Kimura, H., LaCorbiere, M., Vaughan, J., Karr, D., and Fischer, W. H. (1990). Activin is a nerve cell survival molecule. Nature 344, 868–870. Scott, P., Natovitz, P., Coffman, R. L., Pearce, E., and Sher, A. (1988). Immunoregulation of cutaneous leishmaniasis. T cell lines that transfer protective immunity or exacerbation belong to different T helper subsets and respond to distinct parasite antigens. J. Exp. Med. 168, 1675–1684. Semitekolou, M., Alissafi, T., Aggelakopoulou, M., Kourepini, E., Kariyawasam, H. H., Kay, A. B., Robinson, D. S., Lloyd, C. M., Panoutsakopoulou, V., and Xanthou, G. (2009). Activin-A induces regulatory T cells that suppress T helper cell immune responses and protect from allergic airway disease. J. Exp. Med. 206, 1769–1785. Shao, L., Frigon, N. L., Jr., Sehy, D. W., Yu, A. L., Lofgren, J., Schwall, R., and Yu, J. (1992). Regulation of production of activin A in human marrow stromal cells and monocytes. Exp. Hematol. 20, 1235–1242. Shevach, E. M. (2002). CD4þ CD25þ suppressor T cells: More questions than answers. Nat. Rev. Immunol. 2, 389–400. Shull, M. M., Ormsby, I., Kier, A. B., Pawlowski, S., Diebold, R. J., Yin, M., Allen, R., Sidman, C., Proetzel, G., Calvin, D., Annuziata, N., and Doetschman, T. (1992). Targeted disruption of the mouse transforming growth factor-b1 gene results in multifocal inflammatory disease. Nature 359, 693–699. Souchelnytskyi, S., Tamaki, K., Engstrom, U., Wernstedt, C., ten Dijke, P., and Heldin, C. H. (1997). Phosphorylation of Ser465 and Ser467 in the C terminus of Smad2 mediates interaction with Smad4 and is required for transforming growth factor-b signaling. J. Biol. Chem. 272, 28107–28115. Totsuka, Y., Tabuchi, M., Kojima, I., Shibai, H., and Ogata, E. (1988). A novel action of activin A: Stimulation of insulin secretion in rat pancreatic islets. Biochem. Biophys. Res. Commun. 156, 335–339. Tsunawaki, S., Sporn, M., Ding, A., and Nathan, C. (1988). Deactivation of macrophages by transforming growth factor-b. Nature 334, 260–262. Vale, W., Hsueh, A., Rivier, C., and Yu, J. (1990). The inhibin/activin family of hormones and growth factors. In “Peptide Growth Factors and Their Receptors. Handbook of Experimental Pharmacology,” (M. B. Sporn and A. B. Roberts, Eds.), pp. 211–248. Springer-Verlag, New YorkVol. 95. Welgus, H. G., Campbell, E. J., Cury, J. D., Eisen, A. Z., Senior, R. M., Wilhelm, S. M., and Goldberg, G. I. (1990). Neutral metalloproteinases produced by human mononuclear phagocytes. Enzyme profile, regulation, and expression during cellular development. J. Clin. Invest. 86, 1496–1502. Williams, A. O., Knapton, A. D., Geiser, A., Letterio, J. J., and Roberts, A. B. (1996). The liver in transforming growth factor-b1 (TGF-b1) null mutant mice. Ultrastruct. Pathol. 20, 477–490. Woodruff, T. K. (1998). Regulation of cellular and system function by activin. Biochem. Pharmacol. 55, 953–963. Wrana, J. L., Attisano, L., Wieser, R., Ventura, F., and Massague´, J. (1994). Mechanism of activation of the TGF-b receptor. Nature 370, 341–347. Xie, B., Dong, Z., and Fidler, I. J. (1994). Regulatory mechanisms for the expression of type IV collagenases/gelatinases in murine macrophages. J. Immunol. 152, 3637–3644.
Activin in Humoral Immune Responses
253
Yamashita, N., Nakajima, T., Takahashi, H., Kaneoka, H., Mizushima, Y., and Sakane, T. (1993). Effects of activin A on IgE synthesis and cytokine production by human peripheral mononuclear cells. Clin. Exp. Immunol. 94, 214–219. Yamato, K., Koseki, T., Ohguchi, M., Kizaki, M., Ikeda, Y., and Nishihara, T. (1997). Activin A induction of cell-cycle arrest involves modulation of cyclin D2 and p21CIP1/ WAF1 in plasmacytic cells. Mol. Endocrinol. 11, 1044–1052. Yaswen, L., Kulkarni, A. B., Fredrickson, T., Mittleman, B., Schiffman, R., Payne, S., Longenecker, G., Mozes, E., and Karlsson, S. (1996). Autoimmune manifestations in the transforming growth factor-b1 knockout mouse. Blood 87, 1439–1445. Ying, S.-Y. (1988). Inhibins, activins and follistatins: Gonadal proteins modulating the secretion of follicle stimulating hormone. Endocr. Rev. 9, 267–293. Yu, J., Shao, L. E., Frigon, N. J., Jr., Lofgren, J., and Schwall, R. (1996). Induced expression of the new cytokine, activin A, in human monocytes: Inhibition by glucocorticoids and retinoic acid. Immunology 88, 368–374. Zhou, Y., Scamurra, R., Molitor, T. W., and Murtaugh, M. P. (1992). Characterization of transforming growth factor-b1 gene expression in porcine immune cells. Mol. Immunol. 29, 965–970. Zipori, D., and Barda-Saad, M. (2001). Role of activin A in negative regulation of normal and tumor B lymphocytes. J. Leukoc. Biol. 69, 867–873.
C H A P T E R
T H I R T E E N
The Regulation and Functions of Activin and Follistatin in Inflammation and Immunity Mark P. Hedger, Wendy R. Winnall, David J. Phillips,1 and David M. de Kretser2 Contents 256 258 258 260 260 261 262 264 264 265 270 270 275 280 281
I. Introduction II. Molecular Biology of Activin A. Nomenclature, synthesis, and measurement B. The activin A gene C. Other activin subunits D. Receptors and activin signaling E. Follistatin—An activin-binding protein III. Production and Regulation of Activin and Follistatin A. Sites of production and measurement issues B. Regulation during inflammation and immunity IV. Activin Actions A. Activin roles in inflammation, cachexia, and fibrosis B. Activin roles in immunoregulation V. Conclusions References
Abstract The activins are members of the transforming growth factor b superfamily with broad and complex effects on cell growth and differentiation. Activin A has long been known to be a critical regulator of inflammation and immunity, and similar roles are now emerging for activin B, with which it shares 65% sequence homology. These molecules and their binding protein, follistatin, are widely expressed, and their production is increased in many acute and chronic inflammatory conditions. Synthesis and release of the activins are stimulated by Monash Institute of Medical Research, Monash University, Monash Medical Centre, Clayton, Victoria, Australia Current address: Research Services, La Trobe University, Bundoora, Victoria, Australia 2 Current address: Governor of Victoria, Government House, Melbourne, Victoria, Australia 1
Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00013-5
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inflammatory cytokines, Toll-like receptor ligands, and oxidative stress. The activins interact with heterodimeric serine/threonine kinase receptor complexes to activate SMAD transcription factors and the MAP kinase signaling pathways, which mediate inflammation, stress, and immunity. Follistatin binds to the activins with high affinity, thereby obstructing the activin receptor binding site, and targets them to cell surface proteoglycans and lysosomal degradation. Studies on transgenic mice and those with gene knockouts, together with blocking studies using exogenous follistatin, have established that activin A plays critical roles in the onset of cachexia, acute and chronic inflammatory responses such as septicemia, colitis and asthma, and fibrosis. However, activin A also directs the development of monocyte/macrophages, myeloid dendritic cells, and T cell subsets to promote type 2 and regulatory immune responses. The ability of both endogenous and exogenous follistatin to block the proinflammatory and profibrotic actions of activin A has led to interest in this binding protein as a potential therapeutic for limiting the severity of disease and to improve subsequent damage associated with inflammation and fibrosis. However, the ability of activin A to sculpt the subsequent immune response as well means that the full range of effects that might arise from blocking activin bioactivity will need to be considered in any therapeutic applications. ß 2011 Elsevier Inc.
I. Introduction The activins are members of the transforming growth factor b (TGFb) superfamily of growth and differentiation regulating factors that also includes the bone morphogenetic proteins (BMPs), growth differentiation factors (GDFs), nodal and the gonadal hormone, and inhibin. The naming of the activins, their subunits, and their genes was due to their initial identification as regulators of the pituitary hormone, follicle-stimulating hormone (FSH), in antagonism of inhibin (Ling et al., 1986). Inhibin itself comprises one of the activin b-subunits (bA or bB) dimerized to a larger homologous a-subunit (Fig. 13.1) (Stewart et al., 1986). It later became evident that (i) the effects of inhibin on FSH were derived from its ability to antagonize activin produced by the pituitary (Corrigan et al., 1991), and (ii) that activins played regulatory roles in a number of other systems, particularly the hematopoietic and immune system (Broxmeyer et al., 1988; Hedger et al., 1989). Indeed, activin A was isolated several times by several groups, on the basis of its ability to regulate, among others, erythroid differentiation (Eto et al., 1987) and B cell apoptosis (Brosh et al., 1995), resulting in early synonyms for activin such as “erythroid differentiation factor” and “restrictin-P,” respectively. Today, as outlined in this chapter, there is burgeoning interest in the activins because of their ability to regulate inflammation, fibrosis, and immunity in a broad range of systems.
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Figure 13.1 Synthesis and formation of activins and inhibins. Activin A is formed as a homodimer of the inhibin bA-subunit encoded by the INHBA gene, while activin B is formed from inhibin bB-subunits encoded by the INHBB gene. Both homodimers bind to the activin receptor complex to exert a broad range of biological effects on cell development and immunity. The biological activities of activins A and B overlap, but there is also evidence that they possess a slightly different range of activities in different biological systems. Concurrent expression of the inhibin a-subunit, which occurs particularly in steroidogenic tissues like the gonads, adrenals, and placenta, diverts the bA- and bB-subunit to form the heterodimeric inhibins (inhibins A and B). These heterodimers oppose many of the biological actions of the activins, principally by competing for the activin receptor. This means that the a-subunit acts as an intracellular dominant-negative inhibitor of activin, in addition to forming an antagonist that acts at the receptor level. Similar regulatory roles for the bC- and bE-subunits (not shown), which can also heterodimerize with the bA-subunit, and possibly the bB-subunit, have also been suggested. However, the possibility that the inhibins and activins C and E dimers also possess biological activities separate from their ability to interfere with activins A and B cannot yet be excluded.
Although the biology and regulation of the activins has been under investigation for more than 25 years, progress has not been rapid. This is partly related to the complexity of the activin system itself, but also because, as far as inflammation and immunity are concerned, the activins have had a tendency to be overshadowed by the TGFbs, with which they share a crucial signaling pathway (Wrana and Attisano, 2000). It is only more recently that it has become obvious, particularly from studies with knockout mice or experiments employing specific inhibitors, that the activins play
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critical, unique roles in inflammation and immunity that are clearly distinguishable from an ability to act as weak agonists of the TGFbs.
II. Molecular Biology of Activin A. Nomenclature, synthesis, and measurement The canonical representative of the activin family is activin A. Like most members of the TGFb superfamily, activin A is a disulfide-linked dimer of two identical subunits, with additional intra-strand disulfide bonds that form a cysteine knot folding motif (Fig. 13.2) (Greenwald et al., 2004; Harrington et al., 2006; Lin et al., 2006). Activin A is highly conserved across a broad range of species, with 100% conservation between the human, monkey, rat, mouse, bovine, and porcine molecules at the known or predicted subunit protein level, and even 97–98% conservation between the human and predicted reptile, bird, marsupial, and monotreme species sequences. The gene encoding the activin A subunit, named for its location within the activin/inhibin gene subfamily, is called inhibin beta A (INHBA), but more commonly the subunits themselves are designated as the bA-subunits. Activin A is synthesized as a disulfide-linked dimer of 402–426 amino acid precursors, which is processed to the mature bioactive protein of approximately 25 kDa by acidic proteolysis (Fig. 13.2) (Gray and Mason, 1990; Huylebroeck et al., 1990). This mature dimer is the principal circulating form of activin A, and there is no evidence that substantial amounts of activin A remain bound to its prohormone in a latent complex, such as occurs for the TGFbs (Wakefield et al., 1988). Curiously, monomers of the bA-subunit also exist and retain significant residual bioactivity (Hu¨skenHindi et al., 1994; Robertson et al., 1992). However, linkage of the bAsubunit with the inhibin a-subunit during synthesis produces inhibin A, which is a specific inhibitor of activin bioactivity (Fig. 13.1). Expression of the a-subunit is much more restricted than that of the bA-subunit, being largely confined to the gonads, with lower levels expressed in the pituitary, placenta, adrenal gland, and the central nervous system (CNS) (Meunier et al., 1988). The complexity of the molecular processing and multiple potential outcomes from expression of the bA-subunit gene highlight some of the difficulties involved in studying this molecule in vivo. Simple measurement of mRNA levels by RT-PCR, in situ hybridization, or microarray does not provide any information regarding the relative amounts of the potential molecular forms actually present. Immunohistochemistry using subunitspecific antibodies suffers from similar limitations. Bioassays have been developed using cells that show specific responses to activin, such as pituitary cells, B cell plasmacytoma cells, or erythroid cells (Ling et al., 1986;
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Figure 13.2 Activin A synthesis and signaling. Activin A is synthesized as a precursor, which forms disulfide-linked (S–S) homodimers within the cell. Acid hydrolysis cleaves the prohormone sequences (Pro) from the dimerized activin bA-subunits, which spontaneously dissociate and are released from the cell. The activin A dimer binds to type II receptors (ActRII) on the surface of target cells, which oligomerize with type I receptors (ActRI) and phosphorylate specific serine/threonine residues within the intracellular domains of the type I receptors. The type I receptor, in turn, phosphorylates the intracellular proteins SMAD2 and SMAD3, which form heteromeric transcription factor complexes with SMAD4, and translocate into the nucleus to activate target genes. Activin A also induces MAP kinase signaling within cells, leading to downstream effects on proliferation and apoptosis and activation of MAP kinaseresponsive transcription factors.
Phillips et al., 1999; Schwall and Lai, 1991), but these assays are susceptible to interference from other bioactive contaminants. Consequently, most progress has come from assays for dimeric activin A using specific two-site ELISAs (Groome et al., 2001; Knight et al., 1996; Woodruff et al., 1993).
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B. The activin A gene The gene encoding the bA-subunit is located on chromosome 7 in the human and on chromosome 13 in the mouse and consists of three exons and two introns (Tanimoto et al., 1996; Yoshida et al., 1998). Studies on the human, rat, mouse, and bovine 50 regulatory regions have identified a TATA-less core promoter region containing SP1 sites and a proximal enhancer region, which is essential for transcription. This enhancer region consists of at least one AP-1 site, which complexes with JUN family members, and a conserved cAMP-responsive element (CRE)-like site, which fails to complex with the CRE binding protein (CREB) despite the fact that transcription of the bA-subunit can be induced by cAMP (Ardekani et al., 1998; Tanimoto et al., 1996). Instead, this CRE-like site is part of a MARE-like sequence that facilitates transcriptional regulation of the bA-subunit by c-MAF, a transcription factor expressed in Th2 cells (type 2 helper T cells), but not Th1 cells (Ogawa et al., 2006). Transcription of the bA gene is synergistically increased by the combination of c-MAF and NFAT, which binds to a nearby conserved NF-AT consensus site. Transcription of the bA-subunit in response to cAMP is not facilitated by CREB binding, but may instead result from the cAMP-responsive transcription factor AP-2, since several AP-2 binding sites have been identified (Thompson et al., 1994). No consensus NF-kB binding sequences have been identified in the first 3000 nucleotides upstream of the transcription start site in the promoter region of the bA-subunit (Dolter et al., 1998). The presence of five CCAAT-boxes in the proximal promoter region of human bA and three in rodents indicates the possibility of bA transcription regulation by C/EBP family members, which are known to drive upregulation of the expression of a number of cytokines (Cloutier et al., 2009). Human and rodent bA-subunit promoters also contain consensus GATA-binding sites close to their transcription start sites. Although GATA family members have not formally been demonstrated to regulate bA expression, GATA-1 and -4 regulate transcription of the bB-subunit and GATA-1 regulates the inhibin a-subunit in testicular cells (Feng et al., 2000). Conserved CACCC-boxes in the proximal promoter region also indicate a possible regulation by Kru¨ppel-like factors. No splice variants of bA have been described but at least four different length human bA transcripts have been detected, which are predicted to result from differences in the length of the 30 region (Dolter et al., 1998; Tanimoto et al., 1996).
C. Other activin subunits In addition to the bA-subunit and the longer, more distantly related a-subunit, four homologous activin subunits (bB–bE) have been identified (Fang et al., 1996; Ho¨tten et al., 1995; Oda et al., 1995). Activin B is a
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homodimer of the activin bB-subunit gene product, which has approximately 65% sequence similarity to bA (Mason et al., 1985, 1986). Both activin B and the heterodimer activin AB have been identified in biological fluids (Ling et al., 1986; Ludlow et al., 2009). While relatively little is known about the biology of activin AB, activin B not only shares many of the functions of activin A, possibly acting as a slightly weaker activin agonist in many cases, but also may exert functionally distinct effects from those of activin A (Brown et al., 2000, 2003; Corrigan et al., 1991; Mathews and Vale, 1991; Phillips et al., 1999; Thompson et al., 2004). The bB-subunit can also form inhibin by dimerizing with the a-subunit (Fig. 13.1). Activin B expression during inflammation is relatively poorly studied and an ELISA to measure the activin B protein has only recently become available (Ludlow et al., 2009). The bD-subunit gene has only been found in Xenopus (Oda et al., 1995). Expression of the activins C and E subunits is largely, although not exclusively, confined to the liver (Fang et al., 1997; Gold et al., 2004; Ho¨tten et al., 1995; O’Bryan et al., 2000). Discovering biological roles for activins C and E has proven difficult. Whereas mice lacking the bB-subunit have defects of female reproduction and eyelid development (Vassalli et al., 1994) and the lack of bA is neonatal lethal (Matzuk et al., 1995a,b), deletions of bC and bE do not produce obvious phenotypes (Lau et al., 2000). However, the possibility that these subunits may act as competitive inhibitors of activin A or B formation or receptor binding, comparable to the role of the a-subunit, is now emerging from overexpressor models (Gold et al., 2009; Hashimoto et al., 2006; Sekiyama et al., 2009). To date, only the bAand bB-subunits have been shown to be able to dimerize with the a-subunit to form inhibins (Mellor et al., 2000).
D. Receptors and activin signaling Activins A and B act via a classical TGFb family serine/threonine kinase receptor system involving an activin-specific type II receptor (either ActRII or ActRIIB), which binds the ligand and recruits a type I receptor (ActRI; usually activin receptor-like kinase (Alk) 4, but also Alk 2 and Alk 7) (Fig. 13.2) (de Caestecker, 2004; Lin et al., 2006; Tsuchida et al., 2008). Oligomerization of these receptors in the membrane activates serine/threonine kinase activity and phosphorylation of the SMAD proteins 2 and 3, which form a heteromeric complex transcription factor with SMAD 4 (ten Dijke and Hill, 2004; Wrana and Attisano, 2000). This is the same transcription factor cascade induced by the TGFbs acting through their own receptor complexes. SMAD2/3 signaling plays multiple roles in the regulation of inflammation, autoimmunity, and the development of immunoregulatory T cell subsets (Anthoni et al., 2008; Ashcroft et al., 1999; McKarns et al., 2004; Tone et al., 2008; Yang et al., 1999). In common with the
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TGFbs, activin A can also activate alternative signaling pathways involving the MAP kinases (ERK1/2, JNK, and p38 MAPK) (Cocolakis et al., 2001; de Caestecker, 2004; Huang et al., 2006; Ogihara et al., 2003; Zhang et al., 2005a), which are key regulators of inflammation-, stress-, and immunityrelated genes (Agrawal et al., 2003; Guha and Mackman, 2001; Mansell et al., 2004). Regulation of activin action at the receptor level also involves several accessory molecules. Betaglycan is a co-receptor for the activin antagonist inhibin, which promotes a specific interaction between inhibin and the activin type II receptors, and thereby acts as a functional inhibitor of activin binding and signaling (Lewis et al., 2000). BMP and activin membranebound inhibitor (BAMBI) is a transmembrane protein related to the type I receptor proteins, but lacking the intracellular kinase domain, which inhibits activin signaling by binding with the type I and type II receptor subunits and preventing their association and signaling (Onichtchouk et al., 1999). Other membrane-associated proteins that interfere with interactions between activin and/or its receptor subunits include endoglin/CD105 and the GPI-linked membrane protein, Cripto (Barbara et al., 1999; Gray et al., 2003).
E. Follistatin—An activin-binding protein Follistatin, like activin and inhibin, was first identified as a gonadal protein that was able to regulate FSH release from the pituitary (Robertson et al., 1987; Ueno et al., 1987). Further studies established that this was due to the ability of follistatin to bind to the activins with high affinity and neutralize their bioactivity (Nakamura et al., 1990). The follistatin gene comprises five introns and six exons, and alternate splicing produces two mRNA transcripts, one of which retains part of the fifth intronic sequence (Michel et al., 1990; Shimasaki et al., 1988). The retained intron has an early stop codon and produces a proprotein lacking the C-terminal domain that is encoded by the last exon. Subsequent removal of the signal peptide, which is encoded by the first exon, produces two proteins of 288 amino acids (FS288) and 315 amino acids (FS315), respectively (Fig. 13.3). A third protein form of approximately 300–303 amino acids (FS300 or FS303) is produced by proteolytic cleavage of the C-terminal of FS315 (Inouye et al., 1991a; Sugino et al., 1993). This cleaved form of follistatin has only been confirmed in the pig, thus far, but Western blotting studies have described bands which may correspond to this protein in the mouse, rat, bovine, and zebrafish (Hashimoto et al., 1992; Lin et al., 2008; Meinhardt et al., 1998; Ogawa et al., 1997; Wu et al., 2000). Glycosylation of these core proteins produces a number of protein variants ranging in size from 31 to 42 kDa in size. Human follistatin is Asn-glycosylated at two specific sites, but point-
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Figure 13.3 Comparison of the general structures of follistatin and follistatin-related protein. Both follistatin variants are produced from a single gene (FST) by alternate splicing, and comprise an N-terminal domain and three cysteine-rich FS domains of 73–75 amino acids (FS1-3). The activin-binding region of the follistatins is located within the N-terminal region and encompasses the first two FS domains. There is a heparin-binding motif within the FS1 domain, which allows follistatin to bind to both heparin and heparan sulfate-containing proteoglycans on cell surfaces. In contrast to FS288, FS315 has a C-terminal tail extension that normally obstructs the heparinbinding motif. Consequently, FS315 can only bind to proteoglycans after binding of activin causes a conformational change that exposes the heparin-binding motif. A slightly truncated form of the FS315 protein, called FS303, is produced by cleavage of part of the C-terminal. It is generally believed that FS315 and/or FS303 represent the major circulating form of follistatin, and that FS288 tends to be mostly cell associated. Follistatin-related protein (FSRP) is encoded by a separate gene, but possesses a similar structure and the ability to bind to the activins. It does not, however, have the heparinbinding motif that allows facilitates binding to cell surface proteoglycans.
mutation of these sites to remove glycosylation does not change the affinity of FS315 for activin A (Inouye et al., 1991b). Follistatin comprises three homologous 10-cysteine domains and a unique N-terminal, which are involved in the binding of two follistatin molecules to each activin dimer, thereby obstructing the type II receptor binding site (Fig. 13.3; Harrington et al., 2006; Thompson et al., 2005). The binding affinity of activin A and follistatin (50–900 pM) is comparable to the affinity of activin A for the activin receptor itself and is essentially nonreversible (Inouye et al., 1991a; Nakamura et al., 1990; Sugino et al., 1993). The affinity of follistatin binding to activin B is approximately 10-fold lower (Schneyer et al., 2003). FS288 binds intrinsically to heparan sulfate-containing proteoglycans associated with cell surfaces, whereas FS315 can only bind to heparin/heparan sulfate after it binds activin, thereby causing a conformational change in its C-terminus that uncovers the heparin-binding site (Lerch et al., 2007; Sugino et al., 1993). Accordingly, FS315 or its proteolytically cleaved variant is believed to be the main circulating form of
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follistatin. However, administration of heparin, for example, can displace both activin and follistatin, presumably as a complex, from the cell surface into the circulation ( Jones et al., 2004b; Nakamura et al., 1991; Phillips et al., 2000). The follistatin–activin complex bound to cell surface proteoglycans targets activin for removal by a lysosomal degradation pathway (Hashimoto et al., 1997). In addition to activins A and B, follistatin also binds with lower affinity to some other members of the TGFb superfamily, including GDF9, myostatin (GDF8), and BMPs 2, 5, 7, and 8 (Amthor et al., 2004; Cash et al., 2009; Harrington et al., 2006; Iemura et al., 1998). Follistatin does not bind to TGFb1 or TGFb2, but binding of follistatin to TGFb3 has been reported (Nogai et al., 2007). An estimate of the affinity of follistatin for TGFb3 is not yet available. These data highlight the potential for follistatin to modulate the activity of several TGFb family growth factors, particularly at higher concentrations. Moreover, another molecule related to follistatin, called follistatin-related protein (FSRP) or follistatin-like 3 (FSTL3), has been identified (Tsuchida et al., 2000). This protein differs from follistatin in having only two 10-cysteine domains and lacking the heparin-binding sequence (Fig. 13.3), but binds activin A with high affinity and activin B with relatively lower affinity, similar to follistatin (Schneyer et al., 2003). The observation that FSRP/FSTL3 is considerably less effective than follistatin at blocking endogenous activin A in several cell systems (Sidis et al., 2002) suggests that the ability of follistatin to bind to cell surface proteoglycans may be critical for effective inhibition of activin A action. Finally, activin can bind reversibly with several other proteins in biological fluids, such as a2-macroglobulin in the serum, which may act as carriers for activin and control its clearance (Niemuller et al., 1995).
III. Production and Regulation of Activin and Follistatin A. Sites of production and measurement issues Both activins A and B are measurable in the serum (Demura et al., 1992; Knight et al., 1996; Ludlow et al., 2009; McFarlane et al., 1996), but it has proven difficult to identify the main source of these circulating proteins for a number of reasons. First, their genes are very widely expressed. Based on studies in the rat, mouse, pig, and human, the highest levels of bA mRNA expression are found in the ovary, uterus, placenta, male reproductive tract, the CNS, liver, bone marrow, heart, adrenal gland, and fat (Meunier et al., 1988; Schneider et al., 2000; Tuuri et al., 1994) (and our unpublished data). Resting bB mRNA expression is highest in the gonads, placenta, pituitary, uterus, salivary gland, and the CNS (Meunier et al., 1988;
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Schneider et al., 2000; Tuuri et al., 1994). However, it is difficult to quantify how much of this tissue-specific expression is directed to formation of activin A or B dimeric protein in vivo, because of the ability of the bAand bB-subunits to dimerize with the a-subunit or with other b-subunits (de Kretser and Robertson, 1989; Gold et al., 2009; Mellor et al., 2003). Furthermore, follistatin is also expressed in many tissues, particularly the ovary, kidney, uterus, placenta, lymph nodes, thymus, skeletal muscle, CNS, and skin (Hu¨bner et al., 1996; Michel et al., 1990; Schneider et al., 2000; Tuuri et al., 1994) (and our unpublished data) and is present in the circulation (O’Connor et al., 1999). It is rarely possible to discriminate between free activin and follistatin, circulating activin–follistatin complexes, and follistatin or activin–follistatin complexes attached to cell surface proteoglycans, in either biological fluids or tissue extracts using established assays. It is generally assumed that increases in activin protein levels without a corresponding increase in follistatin represent newly synthesized or released activin, but these limitations on measurement represent significant impediments to studies of activin and follistatin biology at present.
B. Regulation during inflammation and immunity 1. Activins Numerous studies have established that activin A increases in the serum and various tissues in acute and chronic inflammatory diseases and models, including inflammatory bowel disease, gastric ulcers, hepatitis and experimental liver damage, rheumatoid arthritis, burns injuries, septicemia, angina and heart failure, meningitis, traumatic brain injury, chronic CNS inflammation, asthma, placental infection, and preeclampsia (Becker et al., 2003; Cho et al., 2003; Ebert et al., 2006; Evron et al., 2007; Gribi et al., 2001; Hardy et al., 2006; Hu¨bner et al., 1997; Hughes et al., 1999; Jones et al., 2007; Karagiannidis et al., 2006; McLean et al., 2008; Michel et al., 1998; Patella et al., 2001; Semitekolou et al., 2009; Tannetta et al., 2003; Yndestad et al., 2004). There is evidence, particularly from patients with septicemia and pulmonary hypertension, that higher serum levels of activin A are related to severity of inflammation and may be predictive of eventual clinical outcomes (Michel et al., 2003; Phillips et al., 2009; Yndestad et al., 2009). Although comparable data for activin B protein are lacking, increases in bB mRNA have been observed in the livers of mice following treatment with lipopolysaccharide (LPS) (our unpublished data), ovalbumin-induced lung inflammation (Rosendahl et al., 2001), experimental colitis (Dohi et al., 2005), and in peripheral blood mononuclear cells in asthma and atopic dermatitis in humans (Wohlfahrt et al., 2003). Moreover, activin A mRNA and protein, as well as bB-subunit mRNA expression, are elevated in fibrosis and during wound healing after trauma or inflammation-induced damage in the skin, liver, kidney, and lungs (Cruise et al., 2004; De Bleser
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et al., 1997; Hu¨bner et al., 1996; Matsuse et al., 1996; McLean et al., 2008; Patella et al., 2006; Wankell et al., 2001a). Based on mRNA expression and protein localization or secretion, activin A is produced by many different cell types during inflammation, immunity, and fibrosis: the most prominent of these are various epithelial cells, hepatocytes and follicular stellate cells in the liver, keratinocytes in the skin, neuronal cells, endothelial cells, smooth muscle cells, bone marrow stromal cells, myeloid cells (monocyte/macrophages and dendritic cells), granulocytes (neutrophils and mast cells), and some T cell subsets (Abe et al., 2002; Cho et al., 2003; De Bleser et al., 1997; Era¨maa et al., 1992; Funaba et al., 2003a; Gribi et al., 2001; Hu¨bner and Werner, 1996; Hughes et al., 1999; Jones et al., 2000b; Matsuse et al., 1996; Ogawa et al., 2006; Okuma et al., 2005a; Patella et al., 2006; Robson et al., 2008; Scutera et al., 2008; Shao et al., 1992; Sugama et al., 2007; Tannetta et al., 2003; Wilson et al., 2006; Yndestad et al., 2009; Yu et al., 1994, 1996). In the inflammatory setting, the expression of the bB-subunit has received relatively very little attention compared with activin A, but there is some evidence that at least some of these cell types also increase production of activin B in response to inflammatory stimuli (Bilezikjian et al., 1998; De Bleser et al., 1997; Sugama et al., 2007). These observations point to important roles for the activins in inflammation and immunity, and in the subsequent tissue repair and fibrotic reactions. In general, regulation of activin A synthesis and release during inflammation remains incompletely characterized. In mice or sheep given a single injection of LPS to induce systemic inflammation, activin A is one of the earliest cytokines to be induced, reaching a peak in the circulation within 1 h, and either preceding or coinciding with the peak levels of the critical proinflammatory cytokine, tumor necrosis factor-a (TNFa) ( Jones et al., 2000a, 2004a, 2007). Curiously, this peak of activin A occurs at least an hour later in rats treated with LPS under similar conditions (O’Bryan et al., 2005). The regulation and source of this early rise are still under investigation. It appears most likely to arise from prestored activin protein or mRNA, rather than from induced expression of the activin gene, which usually takes several hours to respond to LPS in vitro (Era¨maa et al., 1992; Wang et al., 2008; Wilson et al., 2006) (our unpublished data). The rise in circulating activin A also occurs long before any increase in circulating follistatin, indicating that this increase is not due to release of activin– follistatin complexes from cell surface proteoglycans. Activin A remains elevated in the circulation after a single LPS injection for between 5 and 8 h, whereas circulating follistatin begins to increase about 3–4 h after LPS treatment and remains elevated for at least 24 h ( Jones et al., 2000a, 2007; O’Bryan et al., 2005). It is likely that this increase in follistatin is at least partly responsible for the clearance of activin A from the bloodstream.
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Regulation of activin A during inflammation involves the interleukin-1 (IL1)/Toll-like receptor (TLR) signaling pathway. Interleukin-1b (IL1b) is a major early proinflammatory cytokine, although it appears slightly later than TNFa during normal inflammatory responses, and acts via a specific receptor complex, the IL1 receptor (IL1R) (Dinarello, 1996). The TLRs (TLR1-10 in humans, TLR1-13 in rodents) are a major subset of the pattern recognition receptors responsible for recognizing different microbial products or “danger” signals, called pathogen-associated molecular patterns (PAMPs) (Akira and Takeda, 2004; Roach et al., 2005). For example, LPS is a bacterial cell wall product that is recognized by TLR4. Binding of TLR ligands or IL1 to their receptors engages intracellular adapter proteins through a unique Toll/IL1 receptor (TIR) domain in the cytoplasmic tail of the receptor ( Jenkins and Mansell, 2010; O’Neill and Bowie, 2007). Signaling from IL1 and most TLRs (except TLR3) involves parallel and convergent pathways, through the adaptor protein MyD88 and the TNF receptor-associated factors (TRAF) 3 and 6 to activate the prototypical inflammatory transcription factors NF-kB and AP-1. Activation of NF-k B involves phosphorylation and degradation of its repressor subunit IkB, while AP-1 activation occurs via phosphorylation by the MAP kinases, p38 MAPK and JNK (Nakano, 2004). Signaling through MyD88 can also activate the other major MAP kinase, ERK1/2, by a mechanism that is yet to be determined (McNamara et al., 2006). Moreover, TLR3 (the receptor for viral dsRNA) and TLR4 activate a MyD88-independent pathway, through the adaptor TRIF (TIR-domain-containing adapterinducing interferon-b), leading to induction of the transcription factor IRF3 and type 1 interferon (IFNa and b) production, in addition to NFkB (Hertzog et al., 2003). The ability of TLR4 to act via either MyD88 or TRIF is due to its ability to associate with either the Mal (MyD88 adapterlike) or TRAM (TRIF-related adaptor molecule) adaptor proteins, respectively ( Jenkins and Mansell, 2010; O’Neill and Bowie, 2007). Both IL1 and LPS have been shown to be potent stimulators of activin A synthesis and secretion in diverse cell types (Abe et al., 2001, 2002; Bilezikjian et al., 1998; Era¨maa et al., 1992; Hu¨bner and Werner, 1996; Keelan et al., 1998, 2000a; Okuma et al., 2005a,b; Robson et al., 2008; Scutera et al., 2008; Shao et al., 1992; Takahashi et al., 1992; Tannetta et al., 2003; Wang et al., 2008; Wilms et al., 2010; Wilson et al., 2006; Yamashita et al., 1992). The importance of the MyD88 signaling pathway in regulation of activin A secretion was confirmed in vitro using a dominant-negative inactive MyD88 mutant construct and in the LPS-induced systemic inflammation model using MyD88 null mice in vivo ( Jones et al., 2007) (our unpublished data). Regulation of activin A release through activation of other TLRs for both bacterial and viral ligands (specifically TLR2, TLR3, and TLR9) has also been demonstrated in several of these cell types (Ebert et al., 2007; Robson et al., 2008; Scutera et al., 2008; Winnall et al., 2009).
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This has potential clinical relevance, as activin A is upregulated in cultured human airway epithelial cells challenged with human rhinovirus (Leigh et al., 2008), serum activin A levels are increased in chronic viral hepatitis (Patella et al., 2001), and cerebrospinal fluid and serum levels of activin A are increased in both viral and bacterial meningitis (Ebert et al., 2006). Compared with IL1b or TNFa, which are under direct transcriptional control by NF-kB, significant activin A gene expression following IL1/ TLR signaling in vitro occurs several hours later than these early inflammatory cytokines (Era¨maa et al., 1992; Wang et al., 2008; Wilson et al., 2006) (and our unpublished data). This may be due to that fact that the activin A promoter lacks a distinct recognition site for NF-kB binding, although it does contain several AP-1 sites (Ardekani et al., 1998; Tanimoto et al., 1996; Thompson et al., 1994; Yoshida et al., 1998). In human bone marrow stromal cells treated with LPS for 6 h, activin A secretion but not mRNA expression was reduced by inhibition of NF-kB (Scicchitano et al., 2008). However, the upregulation of activin A mRNA levels in HeLa cells infected with Vaccinia virus was able to be reduced by blocking either NF-kB or p38 MAPK (Myskiw et al., 2009). Similarly, the spontaneous production of activin A by porcine cartilage explants in culture could also be blocked by inhibiting NF-kB signaling (Alexander et al., 2007). In a mast cell leukemia cell line, calcium-regulated expression of activin A was able to be partially blocked using inhibitors of either JNK or p38 MAPK (Funaba et al., 2003a). The specific details remain to be fully elucidated, but the balance of published data together with our own unpublished studies using several activin-secreting cell types, including testicular epithelial cells, macrophage cell lines, and neutrophils, suggest that activin A synthesis is most likely regulated through both MyD88-dependent and -independent (TRIF-mediated) signaling pathways via the MAP kinases, and the production of AP-1. Furthermore, the more rapidly responsive regulation of the intracellular storage, translation, and release of activin A, independent of mRNA synthesis, also appears to involve these pathways as well as inflammatory mediators that are themselves under NF-kB and/or AP-1 control. As may be predicted from the presence of multiple phorbol ester responsive elements (i.e., AP-1 and AP-2) in the promoter, activin A gene expression and secretion are also induced by phorbol esters, thereby indicating a stimulatory role for protein kinase C signaling in its regulation (Ardekani et al., 1998; Cho et al., 2003; Dolter et al., 1998; Era¨maa et al., 1992; Hilde´n et al., 1999; Miyanaga et al., 1993; Ogawa et al., 2006; Okuma et al., 2005a; Pawlowski et al., 1997; Shao et al., 1998; Tanimoto et al., 1996; Thompson et al., 1994; Tuuri et al., 1996; Yamashita et al., 1992). Production of activin A by human bone marrow stromal cells in response to LPS has been shown to be reduced by a specific inhibitor of protein kinase C (Scicchitano et al., 2008). Activation of the protein kinase A signaling pathway also increases expression of the bA-subunit in responsive cell
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types, although whether this leads to an increase or decrease in activin A protein secretion may depend upon whether the cells also express the inhibin a-subunit, which is also stimulated through protein kinase A signaling (Ardekani et al., 1998; Keelan et al., 1998; Miyanaga et al., 1993; Okuma et al., 2005a; Tuuri et al., 1996). The relative contributions of these regulatory pathways compared with the MyD88/TRIF/TRAF/MAP kinase pathways in control of activin A during inflammation have not been examined in any detail. In addition to IL1b and TLR ligands such as LPS, several other inflammatory mediators have been shown to regulate activin production in various cell types. The proinflammatory cytokine TNFa, which activates NF-kB and MAP kinase signaling via an alternative MyD88-independent pathway involving TRAF2, stimulates activin A gene expression and protein secretion in bone marrow stromal cells and in fibroblast and keratinocyte cell lines (Dolter et al., 1998; Hu¨bner and Werner, 1996; Shao et al., 1992, 1998; Takahashi et al., 1992), and secretion of activin A by cells of the human placenta and amnion, umbilical vein endothelial cells, peripheral blood mononuclear cells, and monocyte-derived dendritic cells (Keelan et al., 1998, 2000a; Mohan et al., 2001; Scutera et al., 2008; Tannetta et al., 2003). Other cytokines found to stimulate activin A in multiple cell types include granulocyte–monocyte colony stimulating factor, basic fibroblast growth factor, epidermal growth factor, and platelet-derived growth factor (Abe et al., 2002; Alexander et al., 2007; Hu¨bner and Werner, 1996; Mohan et al., 2001; Shao et al., 1992; Uchimaru et al., 1995; Yu et al., 1996). TGFb stimulates activin A mRNA expression and protein secretion in fibroblasts, keratinocytes, and rat hepatic stellate cells, consistent with an interactive role for these TGFb family cytokines in the development of fibrosis (Hu¨bner and Werner, 1996; Ota et al., 2003; Wada et al., 2004; Yamashita et al., 2004). In addition, activin A is stimulated in monocyte/ macrophages and dendritic cells by IFNg and CD40 ligand (CD40L) derived from activated T cells (Abe et al., 2002; Robson et al., 2008; Scutera et al., 2008; Shao et al., 1992; Wilms et al., 2010), and in rat aortic smooth muscle cells by their mitogens, a-thrombin, and angiotensin II (Pawlowski et al., 1997). Conversely, activin A is negatively regulated by glucocorticoids, which are potent anti-inflammatory steroids, in human monocytes, dendritic cells, and bone marrow stromal cells (Scutera et al., 2008; Shao et al., 1998; Yu et al., 1996). Finally, oxidative stress was shown to stimulate activin A release by human placental and endothelial cells, although apparently not by monocytes (Mandang et al., 2007). The mechanisms underlying this last response are unknown, but certainly deserve further attention, given the importance of oxidative stress in inflammation and fibrosis. The regulatory control of activin B in inflammation and immunity has received less attention than has activin A. The regulatory regions of
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the bB-subunit gene share some similarities with those of the bA-subunit (Najmabadi et al., 1993; Thompson et al., 1994), and bB-subunit expression also increases in various models of inflammation and immunity (De Bleser et al., 1997; Dohi et al., 2005; Rosendahl et al., 2001; Sugama et al., 2007; Wohlfahrt et al., 2003). In rat anterior pituitary cells, phorbol esters and IL1b, but not LPS, stimulate expression and synthesis of both bB-subunit mRNA and protein to form activin B dimers (Bilezikjian et al., 1993, 1998). Furthermore, expression of the bB-subunit is increased by phorbol esters, protein kinase A activators, and TGFb in ovarian granulosa cells; although these cells also express high levels of the a-subunit, so much of this production is probably diverted to formation of inhibin rather than activin B under these conditions (Dykema and Mayo, 1994; Era¨maa and Ritvos, 1996; Najmabadi et al., 1993; Turner et al., 1989). 2. Follistatin Follistatin is increased by surgical stress and appears to be a component of the acute phase reaction (Klein et al., 1996a,b). It is elevated in the serum of patients with septicemia (Michel et al., 1998, 2003) and increases in the circulation several hours after activin A appears in LPS-treated mice and sheep ( Jones et al., 2000a, 2004a, 2007; Klein et al., 1996a; Phillips et al., 1998). Similar delayed increases in circulating follistatin levels, as well as increased follistatin mRNA expression in the liver, were observed in mice administered a number of other TLR ligands (our unpublished data). Increased follistatin expression has also been observed in the lungs of patients with pulmonary hypertension (Yndestad et al., 2009), in ovalbumin-induced lung inflammation in mice (Hardy et al., 2006), in the basal epidermal layer of human keloids (Mukhopadhyay et al., 2007), and in the muscle tissue of mice with a genetic model of muscular dystrophy (Abe et al., 2009). However, given that follistatin acts to oppose the actions of the activins, the timing of changes in the expression of follistatin relative to the activins may be a more relevant parameter for consideration. Upregulation of follistatin during inflammation appears to be driven not only by activin itself (Blount et al., 2008; Jones et al., 2007; Wilson et al., 2006) but also by other cytokines, particularly IL1b, TNFa, and IFNg (Abe et al., 2001; Keelan et al., 2000a; Michel et al., 1996; Phillips et al., 1996).
IV. Activin Actions A. Activin roles in inflammation, cachexia, and fibrosis Early studies using an a-subunit knockout mouse model discovered that the resulting increase in systemic activin A levels (and presumably activin B) was accompanied by gonadal tumor development and cachexia, characterized
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by anemia, weight loss, focal necrosis and inflammation of the liver, and atrophy of the stomach (Matzuk et al., 1994). Simultaneously knocking out the ActRII receptor reversed the cachexia and liver pathology in these mice, but not the tumors, indicating that activin signaling plays a role in the development of cachexia (Coerver et al., 1996). Moreover, a significant decrease in body weight has been observed in prepubertal male monkeys administered recombinant activin A for 20 days (Meriggiola et al., 1993). These outcomes also fit with observations that activin inhibits liver growth by stimulating hepatocyte apoptosis (Schwall et al., 1993; Yasuda et al., 1993). Altogether, these data raise the intriguing possibility that activin A may represent the causative ligand in the common pathway to cachexia. Significantly, cachexia involves the induction of critical inflammatory cytokines, specifically TNFa and IL6 and IFNg (Matthys and Billiau, 1997; Tisdale, 1997). Contemporaneous with the early studies implicating activin in cachexia, activin A was demonstrated to stimulate production of a broad range of inflammatory mediators, including IL1b, TNFa, IL6, nitric oxide, prostaglandin E2, and thromboxane, by human, rat, and murine monocyte/ macrophages and/or peripheral blood mononuclear cells (Nu¨sing and Barsig, 1999; Nu¨sing et al., 1995; Yamashita et al., 1993). Subsequently, it was confirmed in macrophage lineage cells that activin induced IkB degradation and translocation of NF-kB into the nucleus (Sugatani et al., 2003), and stimulated ERK1/2 and p38 MAPK signaling (Murase et al., 2001). The precise pathways involved remain poorly defined. Activin A also stimulates the recruitment and maturation of mast cells, which are the key regulatory and effector cells in allergic inflammation (Funaba et al., 2003b, 2006; Murakami et al., 2006). The effects of activin on inflammatory responses appear to be complex (Fig. 13.4). Several studies using a number of different monocyte/macrophage lineage cell types indicate that activin A stimulates inflammatory signaling and inflammatory mediator production in previously untreated cultures, but the effects of activin A become largely inhibitory once the cells are activated. Most notably, activin A was found to inhibit processing of the IL1b precursor into its mature active form in two LPS/phorbol esteractivated human monocyte cell lines (Ohguchi et al., 1998) and inhibit LPS-induced production of nitric oxide, inducible nitric oxide synthase (iNOS), and key proinflammatory cytokines, such as IL1b, TNFa, IL18, and IL6, in rat microglial cells, mouse peritoneal macrophages, and several mouse and human monocyte/macrophage cell lines (Cuschieri et al., 2008; Sugama et al., 2007; Wang et al., 2008, 2009; Wilms et al., 2010; Zhang et al., 2005b; Zhou et al., 2009). Similarly, activin A stimulated baseline cell surface expression of the TLR4 co-receptor CD14, but inhibited TLR4 and CD14 expression in activated macrophages (Wang et al., 2008, 2009; Zhou et al., 2009). Activin A also inhibited foam cell formation, an early event in the onset of atherosclerosis, by regulating scavenger receptor
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Infection, stressor AP-1
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Figure 13.4 Activin and follistatin in control of acute inflammation and fibrosis. Although many of the specific events remain incompletely described, there is considerable evidence that the activins act as critical coregulators or amplifiers of the inflammation and fibrotic response. Activation of inflammatory pathways by microbial molecules, such as LPS, oxidative stress, or other “danger” signals leads to the activation of the proinflammatory transcription factor NF-kB, as well as the production and release of activins A and B, most likely through activation of protein kinase C (PKC) and the p38 and JNK MAP kinases, leading to formation of the JUN family transcription factor, AP-1. The activins, particularly activin A, stimulate the production of NF-kB, as well as a number of inflammatory mediators, including TNFa, IL1b, IL6, iNOS, and (cyclooxygenase 2) COX2. The activins also induce the expression of various profibrotic regulators, which include TNFa, endothelin, and TGFb1. In turn, several of these inflammatory mediators, most notably TNFa, IL1b, and TGFb1, also drive activin production, thereby forming what appears to be a positive feedback loop capable of driving the inflammatory response. Finally, the activins and other inflammatory cytokines, particularly TNFa and IL1b, stimulate the production of follistatin, which acts to block activin, putatively to act as a circuit breaker to limit its ongoing activity. Consequently, introduction of exogenous follistatin during the acute inflammatory phase in experimental animal models is able to reduce the effects of both inflammation and subsequent fibrosis.
expression in cultures of macrophages loaded with acetylated LDL (Kozaki et al., 1997). Furthermore, activin A inhibited many of the downstream actions of IL1 and IL6 on monocytes, lymphocytes, and liver cells (Brosh et al., 1995; Hedger et al., 2000; Russell et al., 1999; Yu et al., 1998). In various tissues associated with human pregnancy, activin A exhibited dose-dependent, biphasic (i.e., negative and positive) effects on inflammatory mediator secretion (Keelan et al., 2000b; Mangioni et al., 2005; Perrier d’Hauterive et al., 2005). Altogether, these data suggest that activin A exerts a proinflammatory effect early in inflammation, particularly at lower doses,
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but once the inflammatory response is ongoing, activin A may act to oppose the inflammatory signaling and inflammatory mediator production. Activin has been implicated as a critical link between the process of inflammation and fibrotic responses (Fig. 13.4). Inflammation generally causes some degree of tissue damage, and fibrosis is an important mechanism underlying the subsequent repair process. However, the replacement of normal tissue with fibrotic tissue can lead to functional impairments, and uncontrolled fibrosis underlies several diseases with serious health consequences, such as idiopathic pulmonary fibrosis, cirrhosis of the liver, and progressive renal failure. Activin A production is stimulated by several known inducers of fibrosis, including TNFa, TGFb1, IL13, endothelin, angiotensin, and thrombin (Dolter et al., 1998; Hardy et al., 2010; Hu¨bner and Werner, 1996; Pawlowski et al., 1997; Reis et al., 2002; Shao et al., 1992, 1998; Takahashi et al., 1992; Wada et al., 2004; Yamashita et al., 2004), and both activin A and bB-subunit expression are increased in many tissues during fibrosis (De Bleser et al., 1997; Gaedeke et al., 2005; Huang et al., 2001; Matsuse et al., 1995, 1996; Mukhopadhyay et al., 2007; Sugiyama et al., 1998; Yamashita et al., 2004). Increasing expression of activin A in the skin of transgenic mice led to excessive dermal fibrosis and increased scarring following skin injury (Munz et al., 1999). Activin A induces expression of several key regulators of the fibrotic process, including TNFa, connective tissue growth factor, type 1 collagen, tissue inhibitor of metalloproteinase-1, plasminogen activator inhibitor 1, and endothelin (Karger et al., 2008; Murakami et al., 2006; Yamashita et al., 2004; Yndestad et al., 2009), and stimulates proliferation of fibroblasts and their differentiation into myofibroblasts, which are crucial steps in the process of fibrosis (Hedger et al., 1989; Karagiannidis et al., 2006; Ohga et al., 1996; Ota et al., 2003; Yamashita et al., 2004). Furthermore, it has been shown that activin A is not only stimulated by TGFb, but activin A also stimulates renal and lung fibroblasts and pancreatic stellate cells to produce TGFb1, providing evidence for a fundamental interdependence of these two cytokines in fibrotic processes (Aoki et al., 2005; Karagiannidis et al., 2006; Ohnishi et al., 2010). Reducing activin bioactivity by administration of exogenous follistatin or increasing follistatin expression using transgenic approaches reduces the severity of inflammatory and fibrotic responses. Administration of follistatin to mice reduced proinflammatory cytokine (TNFa and IL1b) production and reduced mortality in LPS-induced sepsis ( Jones et al., 2007), improved colitis in several models of the disease (Dohi et al., 2005), and inhibited lung mucus production and lymphocyte numbers and cytokine expression in draining mediastinal lymph nodes in an experimental allergic asthma model (Hardy et al., 2006). In rats, follistatin decreased inflammatory injury and fibrosis in models of experimental lung or liver damage (Aoki et al., 2005; Patella et al., 2006). Transgenic overexpression of follistatin in mice also
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reduced the cachexia and liver necrosis/inflammation of inhibin a-subunit deficient mice (Cipriano et al., 2000) and inhibited dermal scar formation after injury (Bamberger et al., 2005; Wankell et al., 2001b). Intriguingly, follistatin blocked the stimulatory actions of TGFb1 on a-smooth muscle actin and/or collagen expression by rat hepatic stellate cells, renal fibroblasts, and lung fibroblasts (Aoki et al., 2005; Wada et al., 2004; Yamashita et al., 2004). This effect of follistatin is almost certainly due to blocking TGFbinduced activin A from these cells, because follistatin does not bind to TGFb1 or block its actions (Harrington et al., 2006; Iemura et al., 1998; Nogai et al., 2007). The observation that expression of follistatin is increased in tissues during fibrosis also suggests a role for endogenous follistatin in controlling the fibrotic effects of activin (Abe et al., 2009; Mukhopadhyay et al., 2007; Wankell et al., 2001a). However, it should be noted that follistatin also binds, albeit less effectively, to myostatin, which is another TGFb family member with established cachectic and profibrotic properties (Amthor et al., 2004; Zimmers et al., 2002). Myostatin is produced by muscle and inhibits muscle growth, acting through either Alk 4 or Alk 5 and the activin type II receptors (Tsuchida et al., 2008). This indicates that myostatin shares a signaling pathway with the activins. The ability of follistatin to prevent both muscle wasting in cachexia and fibrosis in models of muscular dystrophy in mice probably involves interference with myostatin signaling via this pathway, although the relative contributions of endogenous activin and myostatin have not been well characterized in these models (Benny Klimek et al., 2010; Haidet et al., 2008; RodinoKlapac et al., 2009). Novel evidence for the role of the activins in controlling inflammation comes from a recent study showing that overexpressing the bC-subunit in mice causes inflammatory lesions in the liver, which is the tissue where this subunit is normally most highly expressed (Gold et al., 2009). This phenotype was associated with reduced nuclear translocation of SMAD 2. These data may be interpreted in a number of different ways. Increased bC-subunit expression could have interfered with the anti-inflammatory activities of activin A, either by dimerizing with bA-subunits to form activin AC heterodimers and thereby reducing activin A formation, or through the ability of activin AC or C dimers to inhibit activin A receptor binding and/ or signaling (Gold et al., 2009; Muenster et al., 2005). Alternatively, the data may indicate that activin C itself has hitherto unexplored proinflammatory actions in the liver. Liver pathologies were not observed in mice overexpressing the liver-specific bE-subunit (Hashimoto et al., 2006), but expression of the bC- and bE-subunit in the liver was responsive to systemic inflammation in LPS-treated rats (O’Bryan et al., 2000) (our unpublished data). These data point to potential modulatory roles for these so-called inactive activin subunits in inflammation.
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B. Activin roles in immunoregulation Among the earliest roles identified for activin were in control of immune cell development and proliferation in the bone marrow and thymus. Early studies indicated that activin stimulated not only erythrodifferentiation, but also the development of multipotential progenitor cells during hematopoietic development (Broxmeyer et al., 1988; Murata et al., 1988; Yu et al., 1989). Subsequently, it was found that activin A inhibits thymocyte and peripheral T cell activation and proliferation (Hedger and Clarke, 1993; Hedger et al., 1989), and that activin is an important regulator of thymus and thymocyte growth and differentiation (Licona et al., 2006; Rosendahl et al., 2003). Activin modulation of the autocrine/paracrine regulation of T cell growth and differentiation by IL1, IL6, and IFNg has been implicated as a possible mechanism (Hedger et al., 2000; Petraglia et al., 1991). Activin A also has significant inhibitory effects on the survival of normal and transformed B cells (Brosh et al., 1995; Nishihara et al., 1993; Zipori and Barda-Saad, 2001). These data indicate that activin A is a significant immunosuppressive cytokine that exerts direct inhibitory effects on lymphocyte development, activity and survival, which are similar but not entirely identical to the immunosuppressive actions of the TGFbs (Letterio and Roberts, 1998). More recently, however, it has become clear that activin A can also exert more subtle immunoregulatory roles. Initiation of adaptive (antigen-specific) immune responses involves very specific interactions between Th cells and major histocompatibility complex (MHC) class II protein-expressing antigen-presenting cells. Dendritic cells are the most effective antigen-presenting cells, although monocyte/macrophages and even B cells are able to perform this function as well (Banchereau and Steinman, 1998; Sprent and Schaefer, 1989). Antigen-presenting cells process proteins into short antigenic peptides, which are incorporated into a structural groove on the external surface of MHC class II proteins during their assembly, and are recognized by the complementary T cell receptor (TCR) on the appropriate Th cell (Sant et al., 1999; Smith-Garvin et al., 2009). Depending upon the cytokine environment at the site of interaction, this generally leads to either a “type 1” response (Th1 cell), which is characterized by cell-mediated immunity involving cytotoxic T cells, autoimmunity, and rejection responses, or a “type 2” response (Th2 cell) characterized by antibody-mediated immunity and allergy (Constant and Bottomly, 1997; Moser and Murphy, 2000). The principal type 1 cytokines are IFNg and IL12, while the type 2 cytokines include IL4, IL5, and IL13. Depending upon the relative activation status of the antigen-presenting cell and the T cell, the strength of the MHC–TCR interaction and the presence of “immunosuppressive” cytokines such as IL10 and TGFb, this interaction alternatively may produce regulatory and suppressive T cell subsets that may mediate antigen-specific tolerance (Finkelman et al., 1996; Gilliet and Liu, 2002; Sakaguchi et al., 2001; Thompson and Thomas, 2002).
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It has already been noted that activin A is produced by monocyte/ macrophages during inflammation, but production can also be stimulated by interaction with activated T cells and their regulatory molecules, IFNg and CD40L (Abe et al., 2002; Shao et al., 1992; Wilms et al., 2010). Moreover, human dendritic cells produce activin A when activated by LPS or other TLR ligands or by CD40L (Robson et al., 2008; Scutera et al., 2008). However, activin A is also produced by activated murine CD4þ Th2 cells and induces a type 2 macrophage phenotype, characterized by increased arginase-1 and reduced IFNg-induced iNOS and nitric oxide production (Ogawa et al., 2006; Wilms et al., 2010). These observations indicate that activin A may be a type 2 cytokine. Consistent with this suggestion, activin A also stimulates production of IgG, IgA, and IgE by B cells (Lee et al., 2008; Ogawa et al., 2008; Yamashita et al., 1993), while increased activin bA and bB mRNA expression has been associated with a shift toward type 2 macrophage responses in rejecting kidney allografts in mice (Famulski et al., 2008). However, activin A also exerts other immunoregulatory effects. Activin A not only downregulates the inflammatory functions of activated monocyte/macrophages, including production of IL6, an important T and B cell growth factor (Sugama et al., 2007; Wilms et al., 2010), but also can reduce MHC class II expression necessary for antigen-presentation by these cells (Zhou et al., 2009). Activin can also stimulate development of T cells into the effective regulatory T cell subset, CD4þCD25þFoxp3þ Treg cells, in mice (Fig. 13.5) (Huber et al., 2009). Effects of activin A on the immunoregulatory functions of dendritic cells also have been observed. Activin A blocks the inhibitory effects of IL6 on dendritic cell differentiation and promotes the differentiation of human monocytes into myeloid dendritic cells in vitro (Musso et al., 2008; Scutera et al., 2008). Activin A is chemotactic for immature myeloid (monocytederived) dendritic cells from mice and humans (Salogni et al., 2009), and skin dendritic cells (Langerhans cells) are reduced in mice overexpressing follistatin in the skin, although the functions of the Langerhans cells that were recruited appeared to be normal (Stoitzner et al., 2005). In studies of nonsmokers and smokers with or without chronic obstructive pulmonary disease, activin A was expressed in the epithelium, smooth muscle layer, and mononuclear cell infiltrates of the small airways, and the level of expression was found to be correlated with the number of Langerhans-type dendritic cells in these tissues (Van Pottelberge et al., 2010). These studies indicate a positive autocrine and/or paracrine influence of activin A on the recruitment and development of myeloid dendritic cells. Data concerning the role of activin A in the maturation of immature myeloid dendritic cells indicates a more complex suite of effects. Scutera and colleagues found no effect of activin A on LPS-induced maturation of human dendritic cells in culture, or their ability to promote T cell proliferation (Scutera et al., 2008). However, in another study, human immature
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Figure 13.5 Modeling the role of activin in immunoregulation. Based on the data available at present, it appears that activin drives the differentiation of circulating monocytes into either activated macrophages or myeloid dendritic cells, during the initial phase of an immune response. Subsequently, it appears that activin diverts the immune response toward a type 2 or regulatory outcome by stimulating the maturation of activated macrophages into the M2 (alternatively activated) phenotype, thereby reducing their overall production of proinflammatory mediators (TNFa, IL1b, IL6, and iNOS), and diverting Th cells toward either a Th2 or regulatory (Tr) phenotypes through the actions of IL4, IL5, IL13, and IL10. The contributions of other immune cell types involved in this regulation and response, which include NK cells, mast cells, and B cells, have been omitted for clarity, but are outlined elsewhere in this chapter. Consequently, activin is able to promote inflammation and immune activation, but sculpts the subsequent immune response by reducing cell-mediated immunity and promoting fibrosis, antibody-mediated immunity and tolerance.
dendritic cells matured with a cocktail of proinflammatory factors (IL1b, IL6, TNFa, and PGE2) in the presence of activin A displayed a significantly reduced capacity to stimulate T cell proliferation compared with dendritic cells cultured with the inflammatory cocktail alone (Segerer et al., 2008). Blocking activin A activity with follistatin in human dendritic cell cultures did not alter IL6, IL10, or IL12 production stimulated by LPS or intact Escherichia coli, but enhanced the production of IL6, IL10, IL12, TNFa, and several key chemokines in response to CD40L (Robson et al., 2008). A similar effect on IL6 production was observed using siRNA to silence the bA-subunit gene or blocking the Alk 4 receptor with a specific antagonist.
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Moreover, in this same study, follistatin enhanced proliferation, IFNg production, and lytic effector functions of viral antigen-specific CD8þ T cells in cocultures with virally activated dendritic cells (Robson et al., 2008). However, activin A was not able to directly suppress activated CD8þ T cell responses in the absence of dendritic cells. Altogether these data indicate that activin A is able to inhibit the ability of dendritic cells to mature and stimulate T cell activation (Fig. 13.5). This property of activin A is highly significant in the context of immunoregulation, because interaction of T cells with immature dendritic cells tends to produce an immunosuppressive or tolerogenic response ( Jonuleit et al., 2000). Further evidence for the immunoregulatory role of activin A comes from a recent study of interactions between dendritic cells and natural killer (NK) cells (Robson et al., 2009). NK cells are a lymphocyte subset with inherent cytotoxic activities, but are also strong modulators of dendritic cell activity, for example, through production of IFNg (Loza and Perussia, 2001; Raulet, 2004). In cocultures of human NK cells with activated immature myeloid dendritic cells, addition of follistatin considerably enhanced the production of IFNg by the NK cells (Robson et al., 2009). This indicated a role for activin A of dendritic cell origin in the regulation of NK cell production of IFNg, since the NK cells did not produce activin A (Robson et al., 2008). Furthermore, activin A downregulated the production of IFNg, as well as several other proinflammatory cytokines and chemokines, including IL6, TNFa, and IL1b, by NK cells that had been stimulated with IL2 and IL12 (Robson et al., 2009). In contrast, activin A did stimulate IL10 production by these cells, providing further evidence that activin A promotes type 2 or tolerogenic responses. Significantly, activin A appeared to have no effect on the cell killing activities of NK cells, in contrast to TGFb1, which is also a potent suppressor of T cell function (Robson et al., 2009). Asthma is a type 2-associated inflammatory disease. Activin bA-subunit mRNA and protein is increased in CD4þ T cells of moderately asthmatic patients compared with either normal patients or severe asthmatics treated with steroids (Karagiannidis et al., 2006), and both activins A and B were found to be elevated in CD4þ T cells from patients with asthma and atopic dermatitis (another type 2 disease) (Wohlfahrt et al., 2003). In a mouse model of antigen (ovalbumin)-induced allergic asthma, intranasal administration of recombinant mouse follistatin reduced activin A levels in bronchoalveolar lavage fluid (BALF), reduced mucus-producing airway epithelial cells, and decreased the numbers of lymphocytes and proportion of lymphocytes expressing the type 2 cytokines IL4 and IL5 in the draining mediastinal lymph nodes (Hardy et al., 2006). Activin A production in this model is driven by another type 2 cytokine, IL13 (Hardy et al., 2010), and the reduction in disease parameters following follistatin administration was entirely consistent with studies using follistatin to reduce inflammation in other models.
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By contrast, in a similar mouse ovalbumin-induced allergic asthma model, systemic (i.e., intraperitoneal) administration of an activin A neutralizing antibody was found to increase the severity of the disease (Semitekolou et al., 2009). This was accompanied by increased IL4, IL13, and IL10 in the lung and BALF, and increased production of these type 2 cytokines by cells from the draining lymph nodes of the allergic mice. Furthermore, administration of activin A by the same intraperitoneal route suppressed the airway disease parameters and type 2 cytokine responses to an ovalbumin challenge. These observations seem to be counter-intuitive, given that most studies have implicated activin A as a stimulator of type 2 responses. However, detailed examination of the T cells isolated from the draining lymph nodes of the allergic animals treated with activinneutralizing antibody indicated that the anomalous result was due to reduced production of antigen-specific regulatory T cells, which would be capable of suppressing the Th2 effector cell responses responsible for the onset of the disease. Further examination established that activin A was indeed capable of inducing these cell types in vivo and in vitro and that this action of activin A was actually mediated by IL10 and TGFb (Semitekolou et al., 2009). Altogether, these observations were consistent with the hypothesis that circulating activin A protects against chronic airway disease by inducing antigen-specific regulatory T cells that would control both Th1 and Th2 activity, at the local level. Significantly, the same study also reported that local (lung) instillation of activin A had the effect of promoting allergic airway inflammation, which was consistent with the follistatin-treatment model described by Hardy et al. (2006, 2010). This highlights the site-specific nature of activin A effects, whereby local inhibition of activin has a protective effect, presumably by limiting its local inflammatory and immunoregulatory (type 2) actions, but systemic inhibition of activin removes an important control of the overall antigen-specific immune response by regulatory T cells, thereby making the disease potentially worse. In summary, it appears that activin A exerts a complex range of immunoregulatory functions, encompassing effects on (i) the proinflammatory functions of activated monocyte/macrophages; (ii) development and maturation of monocyte-derived dendritic cell; (iii) recruitment, development, and/or survival of T cells, B cells, NK cells, and mast cells; (iv) deviation of local immune responses toward a type 2 phenotype; and (v) production of antigen-specific regulatory T cell subsets (Fig. 13.5). Some of these effects overlap with the activity of TGFb, but other actions appear to be more specific to activin A, thereby identifying this cytokine as a key modulator of immune responses that remains incompletely characterized. Furthermore, the roles of activin B, the other activins and inhibins, and follistatin in these processes continue to emerge and will need be taken into consideration in future studies.
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V. Conclusions In the context of inflammation and immunity, activin A bioactivity is regulated by multiple mechanisms at several different levels. Gene expression, synthesis, and release are induced by bacterial and viral molecular products through activation of TLR, and possibly other pattern recognition receptors, signaling, by various proinflammatory cytokines and other growth factors, and by oxidative stress. Unlike the TGFbs, which are secreted as latent prohormone forms, there is no evidence that activin A remains bound to its precursor sequence after secretion, although it can bind reversibly with several extracellular proteins that may protect it and control its activity, including a2-macroglobulin. However, the intracellular formation of activin A homodimers may be regulated through coexpression of other activin subunits or the a-subunit, leading to formation of heterodimers (i.e., activins AB, AC, AE, and inhibin A) with weak agonist or antagonist properties. The activity of activin A can also be modulated through competitive inhibition by activin homo- and heterodimers and the inhibins or by inhibitory receptor subunit proteins, such as betaglycan, when activin A comes into contact with cells. Most importantly, binding of activin A to follistatin inactivates the molecule in an effectively irreversible manner, since the activin–follistatin complex binds to cell surface heparan sulfate proteoglycans and targets the complex to a lysosomal degradation pathway. This elaborate degree of regulation is believed to be necessary because of the ability of activin to exert potentially deleterious effects on cell survival, differentiation, and function at different sites. In other words, these control mechanisms ensure that activin A (and possibly activin B) acts in a highly localized manner because uncontrolled activin A in the circulation and in tissues could lead to, among other things, liver apoptosis, gastric damage due to parietal cell loss, cachexia, and shock. The role of the activins in inflammation and immunity is far from completely understood. In the early phase of inflammation, activin A promotes the response, and blocking its activity can reduce inflammation and subsequent fibrosis. However, activin A has complex effects on immune cell functions, stimulating inflammatory responses in resting macrophages, but inhibiting the activity of activated macrophages. Activin A also regulates the activity of activated antigen-presenting cells (macrophages and dendritic cells) and T cells, consistent with an important role in maintaining immunoregulation and tolerance. Thus, activin A has both proinflammatory and immunoregulatory functions, which appears to be related to the activation status, duration, and intensity of the immune response. The extent to which the immunoregulatory action of the activins and TGFbs overlap, converge, or interact is an area which requires further investigation. The importance of the activins in controlling immune
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responses in tissues which display immune privilege, such as the gonads, the CNS, and the pregnant uterus, is an area that also merits more attention in the future. In spite of the complexity of the role of the activins in inflammation/ immunity, there is considerable evidence that blocking activin action has a number of therapeutic benefits. The ability of both endogenous and exogenous follistatins to block the proinflammatory and profibrotic actions of activin A has led to interest in this protein as a potential therapeutic for modulating the severity of disease and to improve damage resulting from inflammation and fibrosis. The ability of follistatin to block the actions of myostatin as well as activin has also been explored in muscle disease, such as muscular dystrophy (Rodino-Klapac et al., 2009). More specific inhibitors of activin bioactivity, such as soluble activin receptors, activins C and E, or inhibitors based on activin receptor accessory molecules and prohormone structures may be exploited for similar effect (Benny Klimek et al., 2010; Harrison et al., 2005; Li et al., 2007). Such inhibitors would also have application in liver damage, wasting diseases, and even cancer. However, the ability of activin A to control antigen-specific immune responses as well means that the full range of effects that might arise from blocking activin bioactivity must be considered in any therapeutic applications. This highlights the fact that the route of administration, timing, and doses of agents that interfere with activin actions will be important considerations in the future.
REFERENCES Abe, M., Shintani, Y., Eto, Y., Harada, K., Fujinaka, Y., Kosaka, M., and Matsumoto, T. (2001). Interleukin-1 b enhances and interferon-gamma suppresses activin A actions by reciprocally regulating activin A and follistatin secretion from bone marrow stromal fibroblasts. Clin. Exp. Immunol. 126, 64–68. Abe, M., Shintani, Y., Eto, Y., Harada, K., Kosaka, M., and Matsumoto, T. (2002). Potent induction of activin A secretion from monocytes and bone marrow stromal fibroblasts by cognate interaction with activated T cells. J. Leukoc. Biol. 72, 347–352. Abe, S., Soejima, M., Iwanuma, O., Saka, H., Matsunaga, S., Sakiyama, K., and Ide, Y. (2009). Expression of myostatin and follistatin in Mdx mice, an animal model for muscular dystrophy. Zool. Sci. 26, 315–320. Agrawal, S., Agrawal, A., Doughty, B., Gerwitz, A., Blenis, J., Van Dyke, T., and Pulendran, B. (2003). Cutting edge: Different Toll-like receptor agonists instruct dendritic cells to induce distinct Th responses via differential modulation of extracellular signal-regulated kinase-mitogen-activated protein kinase and c-Fos. J. Immunol. 171, 4984–4989. Akira, S., and Takeda, K. (2004). Toll-like receptor signalling. Nat. Rev. Immunol. 4, 499–511. Alexander, S., Watt, F., Sawaji, Y., Hermansson, M., and Saklatvala, J. (2007). Activin A is an anticatabolic autocrine cytokine in articular cartilage whose production is controlled by fibroblast growth factor 2 and NF-kB. Arthritis Rheum. 56, 3715–3725.
282
Mark P. Hedger et al.
Amthor, H., Nicholas, G., McKinnell, I., Kemp, C. F., Sharma, M., Kambadur, R., and Patel, K. (2004). Follistatin complexes Myostatin and antagonises Myostatin-mediated inhibition of myogenesis. Dev. Biol. 270, 19–30. Anthoni, M., Fyhrquist-Vanni, N., Wolff, H., Alenius, H., and Lauerma, A. (2008). Transforming growth factor-b/Smad3 signalling regulates inflammatory responses in a murine model of contact hypersensitivity. Br. J. Dermatol. 159, 546–554. Aoki, F., Kurabayashi, M., Hasegawa, Y., and Kojima, I. (2005). Attenuation of bleomycininduced pulmonary fibrosis by follistatin. Am. J. Respir. Crit. Care Med. 172, 713–720. Ardekani, A. M., Romanelli, J. C., and Mayo, K. E. (1998). Structure of the rat inhibin and activin bA-subunit gene and regulation in an ovarian granulosa cell line. Endocrinology 139, 3271–3279. Ashcroft, G. S., Yang, X., Glick, A. B., Weinstein, M., Letterio, J. L., Mizel, D. E., Anzano, M., Greenwell-Wild, T., Wahl, S. M., Deng, C., and Roberts, A. B. (1999). Mice lacking Smad3 show accelerated wound healing and an impaired local inflammatory response. Nat. Cell Biol. 1, 260–266. Bamberger, C., Scha¨rer, A., Antsiferova, M., Tychsen, B., Pankow, S., Mu¨ller, M., Ru¨licke, T., Paus, R., and Werner, S. (2005). Activin controls skin morphogenesis and wound repair predominantly via stromal cells and in a concentration-dependent manner via keratinocytes. Am. J. Pathol. 167, 733–747. Banchereau, J., and Steinman, R. M. (1998). Dendritic cells and the control of immunity. Nature 392, 245–252. Barbara, N. P., Wrana, J. L., and Letarte, M. (1999). Endoglin is an accessory protein that interacts with the signaling receptor complex of multiple members of the transforming growth factor-b superfamily. J. Biol. Chem. 274, 584–594. Becker, J. C., Hertel, M., Markmann, A., Shahin, M., Werner, S., Domschke, W., and Pohle, T. (2003). Dynamics and localization of activin A expression in rat gastric ulcers. Scand. J. Gastroenterol. 38, 260–267. Benny Klimek, M. E., Aydogdu, T., Link, M. J., Pons, M., Koniaris, L. G., and Zimmers, T. A. (2010). Acute inhibition of myostatin-family proteins preserves skeletal muscle in mouse models of cancer cachexia. Biochem. Biophys. Res. Commun. 391, 1548–1554. Bilezikjian, L. M., Vaughan, J. M., and Vale, W. W. (1993). Characterization and the regulation of inhibin/activin subunit proteins of cultured rat anterior pituitary cells. Endocrinology 133, 2545–2553. Bilezikjian, L. M., Turnbull, A. V., Corrigan, A. Z., Blount, A. L., Rivier, C. L., and Vale, W. W. (1998). Interleukin-1b regulates pituitary follistatin and inhibin/activin bB mRNA levels and attenuates FSH secretion in response to activin-A. Endocrinology 139, 3361–3364. Blount, A. L., Vaughan, J. M., Vale, W. W., and Bilezikjian, L. M. (2008). A Smad-binding element in intron 1 participates in activin-dependent regulation of the follistatin gene. J. Biol. Chem. 283, 7016–7026. Brosh, N., Sternberg, D., Honigwachs-Sha’anani, J., Lee, B. C., Shav-Tal, Y., Tzehoval, E., Shulman, L. M., Toledo, J., Hacham, Y., Carmi, P., Wen, J., Sasse, J., et al. (1995). The plasmacytoma growth inhibitor restrictin-P is an antagonist of interleukin 6 and interleukin 11. Identification as a stroma-derived activin A. J. Biol. Chem. 270, 29594–29600. Brown, C. W., Houston-Hawkins, D. E., Woodruff, T. K., and Matzuk, M. M. (2000). Insertion of Inhbb into the Inhba locus rescues the Inhba-null phenotype and reveals new activin functions. Nat. Genetics 25, 453–457. Brown, C. W., Li, L., Houston-Hawkins, D. E., and Matzuk, M. M. (2003). Activins are critical modulators of growth and survival. Mol. Endocrinol. 17, 2404–2417. Broxmeyer, H. E., Lu, L., Cooper, S., Schwall, R. H., Mason, A. J., and Nikolics, K. (1988). Selective and indirect modulation of human multipotential and erythroid hematopoietic
Activin and Immunoregulation
283
progenitor cell proliferation by recombinant human activin and inhibin. Proc. Natl. Acad. Sci. USA 85, 9052–9056. Cash, J. N., Rejon, C. A., McPherron, A. C., Bernard, D. J., and Thompson, T. B. (2009). The structure of myostatin:follistatin 288: Insights into receptor utilization and heparin binding. EMBO J. 28, 2662–2676. Cho, S. H., Yao, Z. B., Wang, S. W., Alban, R. F., Barbers, R. G., French, S. W., and Oh, C. K. (2003). Regulation of activin A expression in mast cells and asthma: Its effect on the proliferation of human airway smooth muscle cells. J. Immunol. 170, 4045–4052. Cipriano, S. C., Chen, L., Kumar, T. R., and Matzuk, M. M. (2000). Follistatin is a modulator of gonadal tumor progression and the activin-induced wasting syndrome in inhibin-deficient mice. Endocrinology 141, 2319–2327. Cloutier, A., Guindi, C., Larive´e, P., Dubois, C. M., Amrani, A., and McDonald, P. P. (2009). Inflammatory cytokine production by human neutrophils involves C/EBP transcription factors. J. Immunol. 182, 563–571. Cocolakis, E., Lemay, S., Ali, S., and Lebrun, J. J. (2001). The p38 MAPK pathway is required for cell growth inhibition of human breast cancer cells in response to activin. J. Biol. Chem. 276, 18430–18436. Coerver, K. A., Woodruff, T. K., Finegold, M. J., Mather, J., Bradley, A., and Matzuk, M. M. (1996). Activin signaling through activin receptor type II causes the cachexia-like symptoms in inhibin-deficient mice. Mol. Endocrinol. 10, 534–543. Constant, S. L., and Bottomly, K. (1997). Induction of Th1 and Th2 CD4þ T cell responses: The alternative approaches. Annu. Rev. Immunol. 15, 297–322. Corrigan, A. Z., Bilezikjian, L. M., Carroll, R. S., Bald, L. N., Schmelzer, C. H., Fendly, B. M., Mason, A. J., Chin, W. W., Schwall, R. H., and Vale, W. (1991). Evidence for an autocrine role of activin B within rat anterior pituitary cultures. Endocrinology 128, 1682–1684. Cruise, B. A., Xu, P., and Hall, A. K. (2004). Wounds increase activin in skin and a vasoactive neuropeptide in sensory ganglia. Dev. Biol. 271, 1–10. Cuschieri, J., Bulger, E., Grinsell, R., Garcia, I., and Maier, R. V. (2008). Insulin regulates macrophage activation through activin A. Shock 29, 285–290. De Bleser, P. J., Niki, T., Xu, G., Rogiers, V., and Geerts, A. (1997). Localization and cellular sources of activins in normal and fibrotic rat liver. Hepatology 26, 905–912. de Caestecker, M. (2004). The transforming growth factor-b superfamily of receptors. Cytokine Growth Factor Rev. 15, 1–11. de Kretser, D. M., and Robertson, D. M. (1989). The isolation and physiology of inhibin and related proteins. Biol. Reprod. 40, 33–47. Demura, R., Suzuki, T., Tajima, S., Mitsuhashi, S., Odagiri, E., Eto, Y., Sugino, H., and Demura, H. (1992). Competitive protein binding assay for activin A/EDF using follistatin determination of activin levels in human plasma. Biochem. Biophys. Res. Commun. 185, 1148–1154. Dinarello, C. A. (1996). Biologic basis for interleukin-1 in disease. Blood 87, 2095–2147. Dohi, T., Ejima, C., Kato, R., Kawamura, Y. I., Kawashima, R., Mizutani, N., Tabuchi, Y., and Kojima, I. (2005). Therapeutic potential of follistatin for colonic inflammation in mice. Gastroenterology 128, 411–423. Dolter, K. E., Palyash, J. C., Shao, L. E., and Yu, J. (1998). Analysis of activin A gene expression in human bone marrow stromal cells. J. Cell. Biochem. 70, 8–21. Dykema, J. C., and Mayo, K. E. (1994). Two messenger ribonucleic acids encoding the common bB-chain of inhibin and activin have distinct 50 -initiation sites and are differentially regulated in rat granulosa cells. Endocrinology 135, 702–711. Ebert, S., Phillips, D. J., Jenzewski, P., Nau, R., O0 Connor, A. E., and Michel, U. (2006). Activin A concentrations in human cerebrospinal fluid are age-dependent and elevated in meningitis. J. Neurol. Sci. 250, 50–57.
284
Mark P. Hedger et al.
Ebert, S., Zeretzke, M., Nau, R., and Michel, U. (2007). Microglial cells and peritoneal macrophages release activin A upon stimulation with Toll-like receptor agonists. Neurosci. Lett. 413, 241–244. Era¨maa, M., and Ritvos, O. (1996). Transforming growth factor-beta 1 and -beta 2 induce inhibin and activin beta B-subunit messenger ribonucleic acid levels in cultured human granulosa-luteal cells. Fertil. Steril. 65, 954–960. Era¨maa, M., Hurme, M., Stenman, U. H., and Ritvos, O. (1992). Activin A/erythroid differentiation factor is induced during human monocyte activation. J. Exp. Med. 176, 1449–1452. Eto, Y., Tsuji, T., Takezawa, M., Takano, S., Yokogawa, Y., and Shibai, H. (1987). Purification and characterization of erythroid differentiation factor (EDF) isolated from human leukemia cell line THP-1. Biochem. Biophys. Res. Commun. 142, 1095–1103. Evron, S., Parameswaran, R., Zipori, D., Ezri, T., Sadan, O., and Koren, R. (2007). Activin bA in term placenta and its correlation with placental inflammation in parturients having epidural or systemic meperidine analgesia: A randomized study. J. Clin. Anesth. 19, 168–174. Famulski, K. S., Sis, B., Billesberger, L., and Halloran, P. F. (2008). Interferon-g and donor MHC class I control alternative macrophage activation and activin expression in rejecting kidney allografts: A shift in the Th1–Th2 paradigm. Am. J. Transplant. 8, 547–556. Fang, J., Yin, W., Smiley, E., Wang, S. Q., and Bonadio, J. (1996). Molecular cloning of the mouse activin bE subunit gene. Biochem. Biophys. Res. Commun. 228, 669–674. Fang, J., Wang, S. Q., Smiley, E., and Bonadio, J. (1997). Genes coding for mouse activin bC and bE are closely linked and exhibit a liver-specific expression pattern in adult tissues. Biochem. Biophys. Res. Commun. 231, 655–661. Feng, Z. M., Wu, A. Z., Zhang, Z., and Chen, C. L. (2000). GATA-1 and GATA-4 transactivate inhibin/activin b-B-subunit gene transcription in testicular cells. Mol. Endocrinol. 14, 1820–1835. Finkelman, F. D., Lees, A., Birnbaum, R., Gause, W. C., and Morris, S. C. (1996). Dendritic cells can present antigen in vivo in a tolerogenic or immunogenic fashion. J. Immunol. 157, 1406–1414. Funaba, M., Ikeda, T., Ogawa, K., and Abe, M. (2003a). Calcium-regulated expression of activin A in RBL-2H3 mast cells. Cell. Signal. 15, 605–613. Funaba, M., Ikeda, T., Ogawa, K., Murakami, M., and Abe, M. (2003b). Role of activin A in murine mast cells: Modulation of cell growth, differentiation, and migration. J. Leukoc. Biol. 73, 793–801. Funaba, M., Murakami, M., Ikeda, T., Ogawa, K., Tsuchida, K., and Sugino, H. (2006). Identification of tocopherol-associated protein as an activin/TGF-b-inducible gene in mast cells. Biochim. Biophys. Acta 1763, 900–906. Gaedeke, J., Boehler, T., Budde, K., Neumayer, H. H., and Peters, H. (2005). Glomerular activin A overexpression is linked to fibrosis in anti-Thy1 glomerulonephritis. Nephrol. Dial. Transplant. 20, 319–328. Gilliet, M., and Liu, Y. J. (2002). Generation of human CD8 T regulatory cells by CD40 ligand-activated plasmacytoid dendritic cells. J. Exp. Med. 195, 695–704. Gold, E. J., O0 Bryan, M. K., Mellor, S. L., Cranfield, M., Risbridger, G. P., Groome, N. P., and Fleming, J. S. (2004). Cell-specific expression of bC-activin in the rat reproductive tract, adrenal and liver. Mol. Cell. Endocrinol. 222, 61–69. Gold, E., Jetly, N., O0 Bryan, M. K., Meachem, S., Srinivasan, D., Behuria, S., SanchezPartida, L. G., Woodruff, T., Hedwards, S., Wang, H., McDougall, H., Casey, V., et al. (2009). Activin C antagonizes activin A in vitro and overexpression leads to pathologies in vivo. Am. J. Pathol. 174, 184–195. Gray, A. M., and Mason, A. J. (1990). Requirement for activin A and transforming growth factor-b 1 pro-regions in homodimer assembly. Science 247, 1328–1330.
Activin and Immunoregulation
285
Gray, P. C., Harrison, C. A., and Vale, W. (2003). Cripto forms a complex with activin and type II activin receptors and can block activin signaling. Proc. Natl. Acad. Sci. USA 100, 5193–5198. Greenwald, J., Vega, M. E., Allendorph, G. P., Fischer, W. H., Vale, W., and Choe, S. (2004). A flexible activin explains the membrane-dependent cooperative assembly of TGF-beta family receptors. Mol. Cell 15, 485–489. Gribi, R., Tanaka, T., Harper-Summers, R., and Yu, J. (2001). Expression of activin A in inflammatory arthropathies. Mol. Cell. Endocrinol. 180, 163–167. Groome, N. P., Tsigou, A., Cranfield, M., Knight, P. G., and Robertson, D. M. (2001). Enzyme immunoassays for inhibins, activins and follistatins. Mol. Cell. Endocrinol. 180, 73–77. Guha, M., and Mackman, N. (2001). LPS induction of gene expression in human monocytes. Cell. Signal. 13, 85–94. Haidet, A. M., Rizo, L., Handy, C., Umapathi, P., Eagle, A., Shilling, C., Boue, D., Martin, P. T., Sahenk, Z., Mendell, J. R., and Kaspar, B. K. (2008). Long-term enhancement of skeletal muscle mass and strength by single gene administration of myostatin inhibitors. Proc. Natl. Acad. Sci. USA 105, 4318–4322. Hardy, C. L., O0 Connor, A. E., Yao, J., Sebire, K., de Kretser, D. M., Rolland, J. M., Anderson, G. P., Phillips, D. J., and O0 Hehir, R. E. (2006). Follistatin is a candidate endogenous negative regulator of activin A in experimental allergic asthma. Clin. Exp. Allergy 36, 941–950. Hardy, C. L., Lemasurier, J. S., Olsson, F., Dang, T., Yao, J., Yang, M., Plebanski, M., Phillips, D. J., Mollard, R. A., Rolland, J. M., and O0 Hehir, R. (2010). IL-13 regulates secretion of the TGF-b superfamily cytokine activin A in allergic airway inflammation. Am. J. Respir. Cell Mol. Biol. 42, 667–675. Harrington, A. E., Morris-Triggs, S. A., Ruotolo, B. T., Robinson, C. V., Ohnuma, S., and Hyvo¨nen, M. (2006). Structural basis for the inhibition of activin signalling by follistatin. EMBO J. 25, 1035–1045. Harrison, C. A., Gray, P. C., Vale, W. W., and Robertson, D. M. (2005). Antagonists of activin signaling: Mechanisms and potential biological applications. Trends Endocrinol. Metab. 16, 73–78. Hashimoto, M., Nakamura, T., Inoue, S., Kondo, T., Yamada, R., Eto, Y., Sugino, H., and Muramatsu, M. (1992). Follistatin is a developmentally regulated cytokine in neural differentiation. J. Biol. Chem. 267, 7203–7206. Hashimoto, O., Nakamura, T., Shoji, H., Shimasaki, S., Hayashi, Y., and Sugino, H. (1997). A novel role of follistatin, an activin-binding protein, in the inhibition of activin action in rat pituitary cells. Endocytotic degradation of activin and its acceleration by follistatin associated with cell-surface heparan sulfate. J. Biol. Chem. 272, 13835–13842. Hashimoto, O., Ushiro, Y., Sekiyama, K., Yamaguchi, O., Yoshioka, K., Mutoh, K., and Hasegawa, Y. (2006). Impaired growth of pancreatic exocrine cells in transgenic mice expressing human activin bE subunit. Biochem. Biophys. Res. Commun. 341, 416–424. Hedger, M. P., and Clarke, L. (1993). Isolation of rat blood lymphocytes using a two-step Percoll density gradient. Effect of activin (erythroid differentiation factor) on peripheral T lymphocyte proliferation in vitro. J. Immunol. Methods 163, 133–136. Hedger, M. P., Drummond, A. E., Robertson, D. M., Risbridger, G. P., and de Kretser, D. M. (1989). Inhibin and activin regulate [3H]thymidine uptake by rat thymocytes and 3T3 cells in vitro. Mol. Cell. Endocrinol. 61, 133–138. Hedger, M. P., Phillips, D. J., and de Kretser, D. M. (2000). Divergent cell-specific effects of activin-A on thymocyte proliferation stimulated by phytohemagglutinin, and interleukin 1b or interleukin 6 in vitro. Cytokine 12, 595–602. Hertzog, P. J., O0 Neill, L. A., and Hamilton, J. A. (2003). The interferon in TLR signaling: More than just antiviral. Trends Immunol. 24, 534–539.
286
Mark P. Hedger et al.
Hilde´n, K., Tuuri, T., Era¨maa, M., and Ritvos, O. (1999). Co-ordinate expression of activin A and its type I receptor mRNAs during phorbol ester-induced differentiation of human K562 erythroleukemia cells. Mol. Cell. Endocrinol. 153, 137–145. Ho¨tten, G., Neidhardt, H., Schneider, C., and Pohl, J. (1995). Cloning of a new member of the TGF-b family: A putative new activin bC chain. Biochem. Biophys. Res. Commun. 206, 608–613. Huang, X., Li, D. G., Wang, Z. R., Wei, H. S., Cheng, J. L., Zhan, Y. T., Zhou, X., Xu, Q. F., Li, X., and Lu, H. M. (2001). Expression changes of activin A in the development of hepatic fibrosis. World J. Gastroenterol. 7, 37–41. Huang, H. M., Chiou, H. Y., and Chang, J. L. (2006). Activin A induces erythroid gene expressions and inhibits mitogenic cytokine-mediated K562 colony formation by activating p38 MAPK. J. Cell. Biochem. 98, 789–797. Huber, S., Stahl, F. R., Schrader, J., Lu¨th, S., Presser, K., Carambia, A., Flavell, R. A., Werner, S., Blessing, M., Herkel, J., and Schramm, C. (2009). Activin a promotes the TGF-b-induced conversion of CD4þCD25 T cells into Foxp3þ induced regulatory T cells. J. Immunol. 182, 4633–4640. Hu¨bner, G., and Werner, S. (1996). Serum growth factors and proinflammatory cytokines are potent inducers of activin expression in cultured fibroblasts and keratinocytes. Exp. Cell Res. 228, 106–113. Hu¨bner, G., Hu, Q., Smola, H., and Werner, S. (1996). Strong induction of activin expression after injury suggests an important role of activin in wound repair. Dev. Biol. 173, 490–498. Hu¨bner, G., Brauchle, M., Gregor, M., and Werner, S. (1997). Activin A: A novel player and inflammatory marker in inflammatory bowel disease? Lab. Invest. 77, 311–318. Hughes, P. E., Alexi, T., Walton, M., Williams, C. E., Dragunow, M., Clark, R. G., and Gluckman, P. D. (1999). Activity and injury-dependent expression of inducible transcription factors, growth factors and apoptosis-related genes within the central nervous system. Prog. Neurobiol. 57, 421–450. Hu¨sken-Hindi, P., Tsuchida, K., Park, M., Corrigan, A. Z., Vaughan, J. M., Vale, W. W., and Fischer, W. H. (1994). Monomeric activin A retains high receptor binding affinity but exhibits low biological activity. J. Biol. Chem. 269, 19380–19384. Huylebroeck, D., Van Nimmen, K., Waheed, A., von Figura, K., Marmenout, A., Fransen, L., De Waele, P., Jaspar, J. M., Franchimont, P., and Stunnenberg, H. (1990). Expression and processing of the activin-A/erythroid differentiation factor precursor: A member of the transforming growth factor-b superfamily. Mol. Endocrinol. 4, 1153–1165. Iemura, S., Yamamoto, T. S., Takagi, C., Uchiyama, H., Natsume, T., Shimasaki, S., Sugino, H., and Ueno, N. (1998). Direct binding of follistatin to a complex of bonemorphogenetic protein and its receptor inhibits ventral and epidermal cell fates in early Xenopus embryo. Proc. Natl. Acad. Sci. USA 95, 9337–9342. Inouye, S., Guo, Y., DePaolo, L., Shimonaka, M., Ling, N., and Shimasaki, S. (1991a). Recombinant expression of human follistatin with 315 and 288 amino acids: Chemical and biological comparison with native porcine follistatin. Endocrinology 129, 815–822. Inouye, S., Guo, Y., Ling, N., and Shimasaki, S. (1991b). Site-specific mutagenesis of human follistatin. Biochem. Biophys. Res. Commun. 179, 352–358. Jenkins, K. A., and Mansell, A. (2010). TIR-containing adaptors in Toll-like receptor signalling. Cytokine 49, 237–244. Jones, K. L., Brauman, J. N., Groome, N. P., de Kretser, D. M., and Phillips, D. J. (2000a). Activin A release into the circulation is an early event in systemic inflammation and precedes the release of follistatin. Endocrinology 141, 1905–1908. Jones, R. L., Salamonsen, L. A., Critchley, H. O., Rogers, P. A., Affandi, B., and Findlay, J. K. (2000b). Inhibin and activin subunits are differentially expressed in endometrial cells and leukocytes during the menstrual cycle, in early pregnancy and in women using progestin-only contraception. Mol. Hum. Reprod. 6, 1107–1117.
Activin and Immunoregulation
287
Jones, K. L., de Kretser, D. M., Clarke, I. J., Scheerlinck, J. P., and Phillips, D. J. (2004a). Characterisation of the rapid release of activin A following acute lipopolysaccharide challenge in the ewe. J. Endocrinol. 182, 69–80. Jones, K. L., De Kretser, D. M., and Phillips, D. J. (2004b). Effect of heparin administration to sheep on the release profiles of circulating activin A and follistatin. J. Endocrinol. 181, 307–314. Jones, K. L., Mansell, A., Patella, S., Scott, B. J., Hedger, M. P., de Kretser, D. M., and Phillips, D. J. (2007). Activin A is a critical component of the inflammatory response, and its binding protein, follistatin, reduces mortality in endotoxemia. Proc. Natl. Acad. Sci. USA 104, 16239–16244. Jonuleit, H., Schmitt, E., Schuler, G., Knop, J., and Enk, A. H. (2000). Induction of interleukin 10-producing, nonproliferating CD4þ T cells with regulatory properties by repetitive stimulation with allogeneic immature human dendritic cells. J. Exp. Med. 192, 1213–1222. Karagiannidis, C., Hense, G., Martin, C., Epstein, M., Ru¨ckert, B., Mantel, P. Y., Menz, G., Uhlig, S., Blaser, K., and Schmidt-Weber, C. B. (2006). Activin A is an acute allergen-responsive cytokine and provides a link to TGF-b-mediated airway remodeling in asthma. J. Allergy Clin. Immunol. 117, 111–118. Karger, A., Fitzner, B., Brock, P., Sparmann, G., Emmrich, J., Liebe, S., and Jaster, R. (2008). Molecular insights into connective tissue growth factor action in rat pancreatic stellate cells. Cell. Signal. 20, 1865–1872. Keelan, J. A., Groome, N. P., and Mitchell, M. D. (1998). Regulation of activin-A production by human amnion, decidua and placenta in vitro by pro-inflammatory cytokines. Placenta 19, 429–434. Keelan, J. A., Zhou, R. L., Evans, L. W., Groome, N. P., and Mitchell, M. D. (2000a). Regulation of activin A, inhibin A, and follistatin production in human amnion and choriodecidual explants by inflammatory mediators. J. Soc. Gynecol. Investig. 7, 291–296. Keelan, J. A., Zhou, R. L., and Mitchell, M. D. (2000b). Activin A exerts both pro- and anti-inflammatory effects on human term gestational tissues. Placenta 21, 38–43. Klein, R., Clarke, I. J., Hedger, M. P., and Robertson, D. M. (1996a). Plasma follistatin concentrations increase following lipopolysaccharide administration in sheep. Clin. Exp. Pharmacol. Physiol. 23, 754–755. Klein, R., Robertson, D. M., and Clarke, I. J. (1996b). Studies in sheep examining plasma follistatin elevations due to frequent blood sampling or surgery. Reprod. Fertil. Dev. 8, 273–277. Knight, P. G., Muttukrishna, S., and Groome, N. P. (1996). Development and application of a two-site enzyme immunoassay for the determination of ‘total’ activin-A concentrations in serum and follicular fluid. J. Endocrinol. 148, 267–279. Kozaki, K., Akishita, M., Eto, M., Yoshizumi, M., Toba, K., Inoue, S., Ishikawa, M., Hashimoto, M., Kodama, T., Yamada, N., Orimo, H., and Ouchi, Y. (1997). Role of activin-A and follistatin in foam cell formation of THP-1 macrophages. Arterioscler. Thromb. Vasc. Res. 17, 2389–2394. Lau, A. L., Kumar, T. R., Nishimori, K., Bonadio, J., and Matzuk, M. M. (2000). Activin bC and bE genes are not essential for mouse liver growth, differentiation, and regeneration. Mol. Cell. Biol. 20, 6127–6137. Lee, H. J., Seo, G. Y., Kim, H. A., and Kim, P. H. (2008). Activin A stimulates IgA expression in mouse B cells. Biochem. Biophys. Res. Commun. 366, 574–578. Leigh, R., Oyelusi, W., Wiehler, S., Koetzler, R., Zaheer, R. S., Newton, R., and Proud, D. (2008). Human rhinovirus infection enhances airway epithelial cell production of growth factors involved in airway remodeling. J. Allergy Clin. Immunol. 121(1238– 1245), e1234.
288
Mark P. Hedger et al.
Lerch, T. F., Shimasaki, S., Woodruff, T. K., and Jardetzky, T. S. (2007). Structural and biophysical coupling of heparin and activin binding to follistatin isoform functions. J. Biol. Chem. 282, 15930–15939. Letterio, J. J., and Roberts, A. B. (1998). Regulation of immune responses by TGF-b. Annu. Rev. Immunol. 16, 137–161. Lewis, K. A., Gray, P. C., Blount, A. L., MacConell, L. A., Wiater, E., Bilezikjian, L. M., and Vale, W. (2000). Betaglycan binds inhibin and can mediate functional antagonism of activin signalling. Nature 404, 411–414. Li, Q., Kumar, R., Underwood, K., O’Connor, A. E., Loveland, K. L., Seehra, J. S., and Matzuk, M. M. (2007). Prevention of cachexia-like syndrome development and reduction of tumor progression in inhibin-deficient mice following administration of a chimeric activin receptor type II-murine Fc protein. Mol. Hum. Reprod. 13, 675–683. Licona, P., Chimal-Monroy, J., and Soldevila, G. (2006). Inhibins are the major activin ligands expressed during early thymocyte development. Dev. Dyn. 235, 1124–1132. Lin, S. J., Lerch, T. F., Cook, R. W., Jardetzky, T. S., and Woodruff, T. K. (2006). The structural basis of TGF-b, bone morphogenetic protein, and activin ligand binding. Reproduction 132, 179–190. Lin, S. Y., Craythorn, R. G., O0 Connor, A. E., Matzuk, M. M., Girling, J. E., Morrison, J. R., and de Kretser, D. M. (2008). Female infertility and disrupted angiogenesis are actions of specific follistatin isoforms. Mol. Endocrinol. 22, 415–429. Ling, N., Ying, S. Y., Ueno, N., Shimasaki, S., Esch, F., Hotta, M., and Guillemin, R. (1986). Pituitary FSH is released by a heterodimer of the b-subunits from the two forms of inhibin. Nature 321, 779–782. Loza, M. J., and Perussia, B. (2001). Final steps of natural killer cell maturation: A model for type 1-type 2 differentiation? Nat. Immunol. 2, 917–924. Ludlow, H., Phillips, D. J., Myers, M., McLachlan, R. I., de Kretser, D. M., Allan, C. A., Anderson, R. A., Groome, N. P., Hyvonen, M., Colin Duncan, W., and Muttukrishna, S. (2009). A new ‘total’ activin B enzyme-linked immunosorbent assay (ELISA): Development and validation for human samples. Clin. Endocrinol. 71, 867–873. Mandang, S., Manuelpillai, U., and Wallace, E. M. (2007). Oxidative stress increases placental and endothelial cell activin A secretion. J. Endocrinol. 192, 485–493. Mangioni, S., Vigano, P., Florio, P., Borghi, O., Vignali, M., Petraglia, F., and Di Blasio, A. M. (2005). Effect of activin A on tumor necrosis factor-a/intercellular adhesion molecule-1 pathway in endometrial stromal cells. Eur. J. Obstet. Gynecol. Reprod. Biol. 123, 218–223. Mansell, A., Brint, E., Gould, J. A., O0 Neill, L. A., and Hertzog, P. J. (2004). Mal interacts with tumor necrosis factor receptor-associated factor (TRAF)-6 to mediate NF-kappaB activation by toll-like receptor (TLR)-2 and TLR4. J. Biol. Chem. 279, 37227–37230. Mason, A. J., Hayflick, J. S., Ling, N., Esch, F., Ueno, N., Ying, S. Y., Guillemin, R., Niall, H., and Seeburg, P. H. (1985). Complementary DNA sequences of ovarian follicular fluid inhibin show precursor structure and homology with transforming growth factor-b. Nature 318, 659–663. Mason, A. J., Niall, H. D., and Seeburg, P. H. (1986). Structure of two human ovarian inhibins. Biochem. Biophys. Res. Commun. 135, 957–964. Mathews, L. S., and Vale, W. W. (1991). Expression cloning of an activin receptor, a predicted transmembrane serine kinase. Cell 65, 973–982. Matsuse, T., Fukuchi, Y., Eto, Y., Matsui, H., Hosoi, T., Oka, T., Ohga, E., Nagase, T., and Orimo, H. (1995). Expression of immunoreactive and bioactive activin A protein in adult murine lung after bleomycin treatment. Am. J. Respir. Cell Mol. Biol. 13, 17–24. Matsuse, T., Ikegami, A., Ohga, E., Hosoi, T., Oka, T., Kida, K., Fukayama, M., Inoue, S., Nagase, T., Ouchi, Y., and Fukuchi, Y. (1996). Expression of immunoreactive activin A protein in remodeling lesions associated with interstitial pulmonary fibrosis. Am. J. Pathol. 148, 707–713.
Activin and Immunoregulation
289
Matthys, P., and Billiau, A. (1997). Cytokines and cachexia. Nutrition 13, 763–770. Matzuk, M. M., Finegold, M. J., Mather, J. P., Krummen, L., Lu, H., and Bradley, A. (1994). Development of cancer cachexia-like syndrome and adrenal tumors in inhibindeficient mice. Proc. Natl. Acad. Sci. USA 91, 8817–8821. Matzuk, M. M., Kumar, T. R., and Bradley, A. (1995a). Different phenotypes for mice deficient in either activins or activin receptor type II. Nature 374, 356–360. Matzuk, M. M., Kumar, T. R., Vassalli, A., Bickenbach, J. R., Roop, D. R., Jaenisch, R., and Bradley, A. (1995b). Functional analysis of activins during mammalian development. Nature 374, 354–356. McFarlane, J. R., Foulds, L. M., Pisciotta, A., Robertson, D. M., and de Kretser, D. M. (1996). Measurement of activin in biological fluids by radioimmunoassay, utilizing dissociating agents to remove the interference of follistatin. Eur. J. Endocrinol. 134, 481–489. McKarns, S. C., Schwartz, R. H., and Kaminski, N. E. (2004). Smad3 is essential for TGF-b1 to suppress IL-2 production and TCR-induced proliferation, but not IL-2-induced proliferation. J. Immunol. 172, 4275–4284. McLean, C. A., Cleland, H., Moncrieff, N. J., Barton, R. J., de Kretser, D. M., and Phillips, D. J. (2008). Temporal expression of activin in acute burn wounds–From inflammatory cells to fibroblasts. Burns 34, 50–55. McNamara, N., Gallup, M., Sucher, A., Maltseva, I., McKemy, D., and Basbaum, C. (2006). AsialoGM1 and TLR5 cooperate in flagellin-induced nucleotide signaling to activate Erk1/2. Am. J. Respir. Cell Mol. Biol. 34, 653–660. Meinhardt, A., O0 Bryan, M. K., McFarlane, J. R., Loveland, K. L., Mallidis, C., Foulds, L. M., Phillips, D. J., and de Kretser, D. M. (1998). Localization of follistatin in the rat testis. J. Reprod. Fertil. 112, 233–241. Mellor, S. L., Cranfield, M., Ries, R., Pedersen, J., Cancilla, B., de Kretser, D., Groome, N. P., Mason, A. J., and Risbridger, G. P. (2000). Localization of activin bA-, bB-, and bC-subunits in humanprostate and evidence for formation of new activin heterodimers of bC-subunit. J. Clin. Endocrinol. Metab. 85, 4851–4858. Mellor, S. L., Ball, E. M., O0 Connor, A. E., Ethier, J. F., Cranfield, M., Schmitt, J. F., Phillips, D. J., Groome, N. P., and Risbridger, G. P. (2003). Activin bC-subunit heterodimers provide a new mechanism of regulating activin levels in the prostate. Endocrinology 144, 4410–4419. Meriggiola, M. C., Paulsen, C. A., and Bremner, W. J. (1993). Long term human recombinant activin administration to prepubertal male monkeys. In Proceedings of the 75th Annual Meeting of the Endocrine Society (p. 487, Las Vegas, Nevada). Meunier, H., Rivier, C., Evans, R. M., and Vale, W. (1988). Gonadal and extragonadal expression of inhibin a, bA, and bB subunits in various tissues predicts diverse functions. Proc. Natl. Acad. Sci. USA 85, 247–251. Michel, U., Albiston, A., and Findlay, J. K. (1990). Rat follistatin: Gonadal and extragonadal expression and evidence for alternative splicing. Biochem. Biophys. Res. Commun. 173, 401–407. Michel, U., Schneider, O., Kirchhof, C., Meisel, S., Smirnov, A., Wiltfang, J., and Rieckmann, P. (1996). Production of follistatin in porcine endothelial cells: Differential regulation by bacterial compounds and the synthetic glucocorticoid RU 28362. Endocrinology 137, 4925–4934. Michel, U., Shintani, Y., and Nau, R. (1998). Serum follistatin concentrations are increased in patients with septicaemia. Clin. Endocrinol. 48, 413–417. Michel, U., Ebert, S., Phillips, D., and Nau, R. (2003). Serum concentrations of activin and follistatin are elevated and run in parallel in patients with septicemia. Eur. J. Endocrinol. 148, 559–564.
290
Mark P. Hedger et al.
Miyanaga, K., Erickson, G. F., DePaolo, L. V., Ling, N., and Shimasaki, S. (1993). Differential control of activin, inhibin and follistatin proteins in cultured rat granulosa cells. Biochem. Biophys. Res. Commun. 194, 253–258. Mohan, A., Asselin, J., Sargent, I. L., Groome, N. P., and Muttukrishna, S. (2001). Effect of cytokines and growth factors on the secretion of inhibin A, activin A and follistatin by term placental villous trophoblasts in culture. Eur. J. Endocrinol. 145, 505–511. Moser, M., and Murphy, K. M. (2000). Dendritic cell regulation of TH1–TH2 development. Nat. Immunol. 1, 199–205. Muenster, U., Harrison, C. A., Donaldson, C., Vale, W., and Fischer, W. H. (2005). An activin-A/C chimera exhibits activin and myostatin antagonistic properties. J. Biol. Chem. 280, 36626–36632. Mukhopadhyay, A., Chan, S. Y., Lim, I. J., Phillips, D. J., and Phan, T. T. (2007). The role of the activin system in keloid pathogenesis. Am. J. Physiol. Cell Physiol. 292, C1331–C1338. Munz, B., Smola, H., Engelhardt, F., Bleuel, K., Brauchle, M., Lein, I., Evans, L. W., Huylebroeck, D., Balling, R., and Werner, S. (1999). Overexpression of activin A in the skin of transgenic mice reveals new activities of activin in epidermal morphogenesis, dermal fibrosis and wound repair. EMBO J. 18, 5205–5215. Murakami, M., Ikeda, T., Saito, T., Ogawa, K., Nishino, Y., Nakaya, K., and Funaba, M. (2006). Transcriptional regulation of plasminogen activator inhibitor-1 by transforming growth factor-b, activin A and microphthalmia-associated transcription factor. Cell. Signal. 18, 256–265. Murase, Y., Okahashi, N., Koseki, T., Itoh, K., Udagawa, N., Hashimoto, O., Sugino, H., Noguchi, T., and Nishihara, T. (2001). Possible involvement of protein kinases and Smad2 signaling pathways on osteoclast differentiation enhanced by activin A. J. Cell. Physiol. 188, 236–242. Murata, M., Eto, Y., Shibai, H., Sakai, M., and Muramatsu, M. (1988). Erythroid differentiation factor is encoded by the same mRNA as that of the inhibin bAchain. Proc. Natl. Acad. Sci. USA 85, 2434–2438. Musso, T., Scutera, S., Vermi, W., Daniele, R., Fornaro, M., Castagnoli, C., Alotto, D., Ravanini, M., Cambieri, I., Salogni, L., Elia, A. R., Giovarelli, M., et al. (2008). Activin A induces Langerhans cell differentiation in vitro and in human skin explants. PLoS ONE 3, e3271. Myskiw, C., Arsenio, J., van Bruggen, R., Deschambault, Y., and Cao, J. (2009). Vaccinia virus E3 suppresses expression of diverse cytokines through inhibition of the PKR, NF-kB, and IRF3 pathways. J. Virol. 83, 6757–6768. Najmabadi, H., Rosenberg, L. A., Yuan, Q. X., Reyaz, G., and Bhasin, S. (1993). Transcriptional regulation of inhibin bB messenger ribonucleic acid levels in TM.4 or primary rat Sertoli cells by 8-bromo-cyclic adenosine monophosphate. Mol. Endocrinol. 7, 561–569. Nakamura, T., Takio, K., Eto, Y., Shibai, H., Titani, K., and Sugino, H. (1990). Activinbinding protein from rat ovary is follistatin. Science 247, 836–838. Nakamura, T., Sugino, K., Titani, K., and Sugino, H. (1991). Follistatin, an activin-binding protein, associates with heparan sulfate chains of proteoglycans on follicular granulosa cells. J. Biol. Chem. 266, 19432–19437. Nakano, H. (2004). Signaling crosstalk between NF-kB and JNK. Trends Immunol. 25, 402–405. Niemuller, C. A., Randall, K. J., Webb, D. J., Gonias, S. L., and LaMarre, J. (1995). a2macroglobulin conformation determines binding affinity for activin A and plasma clearance of activin A/a2-macroglobulin complex. Endocrinology 136, 5343–5349. Nishihara, T., Okahashi, N., and Ueda, N. (1993). Activin A induces apoptotic cell death. Biochem. Biophys. Res. Commun. 197, 985–991.
Activin and Immunoregulation
291
Nogai, H., Rosowski, M., Grun, J., Rietz, A., Debus, N., Schmidt, G., Lauster, C., Janitz, M., Vortkamp, A., and Lauster, R. (2007). Follistatin antagonizes transforming growth factor-b3-induced epithelial-mesenchymal transition in vitro: Implications for murine palatal development supported by microarray analysis. Differentiation 76, 404–416. Nu¨sing, R. M., and Barsig, J. (1999). Induction of prostanoid, nitric oxide, and cytokine formation in rat bone marrow derived macrophages by activin A. Br. J. Pharmacol. 127, 919–926. Nu¨sing, R. M., Mohr, S., and Ullrich, V. (1995). Activin A and retinoic acid synergize in cyclooxygenase-1 and thromboxane synthase induction during differentiation of J774.1 macrophages. Eur. J. Biochem. 227, 130–136. O0 Bryan, M. K., Sebire, K. L., Gerdprasert, O., Hedger, M. P., Hearn, M. T., and de Kretser, D. M. (2000). Cloning and regulation of the rat activin bE subunit. J. Mol. Endocrinol. 24, 409–418. O0 Bryan, M. K., Gerdprasert, O., Nikolic-Paterson, D. J., Meinhardt, A., Muir, J. A., Foulds, L. M., Phillips, D. J., de Kretser, D. M., and Hedger, M. P. (2005). Cytokine profiles in the testes of rats treated with lipopolysaccharide reveal localized suppression of inflammatory responses. Am. J. Physiol. Regul. Integr. Comp. Physiol. 288, R1744–R1755. O0 Connor, A. E., McFarlane, J. R., Hayward, S., Yohkaichiya, T., Groome, N. P., and de Kretser, D. M. (1999). Serum activin A and follistatin concentrations during human pregnancy: A cross-sectional and longitudinal study. Hum. Reprod. 14, 827–832. O0 Neill, L. A., and Bowie, A. G. (2007). The family of five: TIR-domain-containing adaptors in Toll-like receptor signalling. Nat. Rev. Immunol. 7, 353–364. Oda, S., Nishimatsu, S., Murakami, K., and Ueno, N. (1995). Molecular cloning and functional analysis of a new activin b subunit: A dorsal mesoderm-inducing activity in Xenopus. Biochem. Biophys. Res. Commun. 210, 581–588. Ogawa, K., Hashimoto, O., Kurohmaru, M., Mizutani, T., Sugino, H., and Hayashi, Y. (1997). Follistatin-like immunoreactivity in the cytoplasm and nucleus of spermatogenic cells in the rat. Eur. J. Endocrinol. 137, 523–529. Ogawa, K., Funaba, M., Chen, Y., and Tsujimoto, M. (2006). Activin A functions as a Th2 cytokine in the promotion of the alternative activation of macrophages. J. Immunol. 177, 6787–6794. Ogawa, K., Funaba, M., and Tsujimoto, M. (2008). A dual role of activin A in regulating immunoglobulin production of B cells. J. Leukoc. Biol. 83, 1451–1458. Ogihara, T., Watada, H., Kanno, R., Ikeda, F., Nomiyama, T., Tanaka, Y., Nakao, A., German, M. S., Kojima, I., and Kawamori, R. (2003). p38 MAPK is involved in activin A- and hepatocyte growth factor-mediated expression of pro-endocrine gene neurogenin 3 in AR42J-B13 cells. J. Biol. Chem. 278, 21693–21700. Ohga, E., Matsuse, T., Teramoto, S., Katayama, H., Nagase, T., Fukuchi, Y., and Ouchi, Y. (1996). Effects of activin A on proliferation and differentiation of human lung fibroblasts. Biochem. Biophys. Res. Commun. 228, 391–396. Ohguchi, M., Yamato, K., Ishihara, Y., Koide, M., Ueda, N., Okahashi, N., Noguchi, T., Kizaki, M., Ikeda, Y., Sugino, H., and Nisihara, T. (1998). Activin A regulates the production of mature interleukin-1b and interleukin-1 receptor antagonist in human monocytic cells. J. Interferon Cytokine Res. 18, 491–498. Ohnishi, N., Miyata, T., Ohnishi, H., Yasuda, H., Tamada, K., Ueda, N., Mashita, H., and Suganao, K. (2010). Activin A is an autocrine activator of rat pancreatic stellate cells: Potential therapeutic role of follistatin for pancreatic fibrosis. Gut 52, 1487–1493. Okuma, Y., O0 Connor, A. E., Muir, J. A., Stanton, P. G., de Kretser, D. M., and Hedger, M. P. (2005a). Regulation of activin A and inhibin B secretion by inflammatory mediators in adult rat Sertoli cell cultures. J. Endocrinol. 187, 125–134.
292
Mark P. Hedger et al.
Okuma, Y., Saito, K., O0 Connor, A. E., Phillips, D. J., de Kretser, D. M., and Hedger, M. P. (2005b). Reciprocal regulation of activin A and inhibin B by interleukin-1 (IL-1) and follicle-stimulating hormone (FSH) in rat Sertoli cells in vitro. J. Endocrinol. 185, 99–110. Onichtchouk, D., Chen, Y. G., Dosch, R., Gawantka, V., Delius, H., Massague, J., and Niehrs, C. (1999). Silencing of TGF-beta signalling by the pseudoreceptor BAMBI. Nature 401, 480–485. Ota, F., Maeshima, A., Yamashita, S., Ikeuchi, H., Kaneko, Y., Kuroiwa, T., Hiromura, K., Ueki, K., Kojima, I., and Nojima, Y. (2003). Activin A induces cell proliferation of fibroblast-like synoviocytes in rheumatoid arthritis. Arthritis Rheum. 48, 2442–2449. Patella, S., Phillips, D. J., de Kretser, D. M., Evans, L. W., Groome, N. P., and Sievert, W. (2001). Characterization of serum activin-A and follistatin and their relation to virological and histological determinants in chronic viral hepatitis. J. Hepatol. 34, 576–583. Patella, S., Phillips, D. J., Tchongue, J., de Kretser, D. M., and Sievert, W. (2006). Follistatin attenuates early liver fibrosis: Effects on hepatic stellate cell activation and hepatocyte apoptosis. Am. J. Physiol. Gastrointest. Liver Physiol. 290, G137–G144. Pawlowski, J. E., Taylor, D. S., Valentine, M., Hail, M. E., Ferrer, P., Kowala, M. C., and Molloy, C. J. (1997). Stimulation of activin A expression in rat aortic smooth muscle cells by thrombin and angiotensin II correlates with neointimal formation in vivo. J. Clin. Invest. 100, 639–648. Perrier d0 Hauterive, S., Charlet-Renard, C., Dubois, M., Berndt, S., Goffin, F., Foidart, J. M., and Geenen, V. (2005). Human endometrial leukemia inhibitory factor and interleukin-6: Control of secretion by transforming growth factor-b-related members. Neuroimmunomodulation 12, 157–163. Petraglia, F., Sacerdote, P., Cossarizza, A., Angioni, S., Genazzani, A. D., Franceschi, C., Muscettola, M., and Grasso, G. (1991). Inhibin and activin modulate human monocyte chemotaxis and human lymphocyte interferon-g production. J. Clin. Endocrinol. Metab. 72, 496–502. Phillips, D. J., Hedger, M. P., McFarlane, J. R., Klein, R., Clarke, I. J., Tilbrook, A. J., Nash, A. D., and de Kretser, D. M. (1996). Follistatin concentrations in male sheep increase following sham castration/castration or injection of interleukin-1b. J. Endocrinol. 151, 119–124. Phillips, D. J., de Kretser, D. M., Pfeffer, A., Chie, W. N., and Moore, L. G. (1998). Follistatin has a biphasic response but follicle-stimulating hormone is unchanged during an inflammatory episode in growing lambs. J. Endocrinol. 156, 77–82. Phillips, D. J., Brauman, J. N., Mason, A. J., de Kretser, D. M., and Hedger, M. P. (1999). A sensitive and specific in vitro bioassay for activin using a mouse plasmacytoma cell line, MPC-11. J. Endocrinol. 162, 111–116. Phillips, D. J., Jones, K. L., McGaw, D. J., Groome, N. P., Smolich, J. J., Parsson, H., and de Kretser, D. M. (2000). Release of activin and follistatin during cardiovascular procedures is largely due to heparin administration. J. Clin. Endocrinol. Metab. 85, 2411–2415. Phillips, D. J., de Kretser, D. M., and Hedger, M. P. (2009). Activin and related proteins in inflammation: Not just interested bystanders. Cytokine Growth Factor Rev. 20, 153–164. Raulet, D. H. (2004). Interplay of natural killer cells and their receptors with the adaptive immune response. Nat. Immunol. 5, 996–1002. Reis, F. M., Luisi, S., Florio, P., Degrassi, A., and Petraglia, F. (2002). Corticotropinreleasing factor, urocortin and endothelin-1 stimulate activin A release from cultured human placental cells. Placenta 23, 522–525. Roach, J. C., Glusman, G., Rowen, L., Kaur, A., Purcell, M. K., Smith, K. D., Hood, L. E., and Aderem, A. (2005). The evolution of vertebrate Toll-like receptors. Proc. Natl. Acad. Sci. USA 102, 9577–9582.
Activin and Immunoregulation
293
Robertson, D. M., Klein, R., de Vos, F. L., McLachlan, R. I., Wettenhall, R. E., Hearn, M. T., Burger, H. G., and de Kretser, D. M. (1987). The isolation of polypeptides with FSH suppressing activity from bovine follicular fluid which are structurally different to inhibin. Biochem. Biophys. Res. Commun. 149, 744–749. Robertson, D. M., Foulds, L. M., Prisk, M., and Hedger, M. P. (1992). Inhibin/activin b-subunit monomer: Isolation and characterization. Endocrinology 130, 1680–1687. Robson, N. C., Phillips, D. J., McAlpine, T., Shin, A., Svobodova, S., Toy, T., Pillay, V., Kirkpatrick, N., Zanker, D., Wilson, K., Helling, I., Wei, H., et al. (2008). Activin-A: A novel dendritic cell-derived cytokine that potently attenuates CD40 ligand-specific cytokine and chemokine production. Blood 111, 2733–2743. Robson, N. C., Wei, H., McAlpine, T., Kirkpatrick, N., Cebon, J., and Maraskovsky, E. (2009). Activin-A attenuates several human natural killer cell functions. Blood 113, 3218–3225. Rodino-Klapac, L. R., Haidet, A. M., Kota, J., Handy, C., Kaspar, B. K., and Mendell, J. R. (2009). Inhibition of myostatin with emphasis on follistatin as a therapy for muscle disease. Muscle Nerve 39, 283–296. Rosendahl, A., Checchin, D., Fehniger, T. E., ten Dijke, P., Heldin, C. H., and Sideras, P. (2001). Activation of the TGF-b/activin-Smad2 pathway during allergic airway inflammation. Am. J. Respir. Cell Mol. Biol. 25, 60–68. Rosendahl, A., Speletas, M., Leandersson, K., Ivars, F., and Sideras, P. (2003). Transforming growth factor-b- and Activin-Smad signaling pathways are activated at distinct maturation stages of the thymopoeisis. Int. Immunol. 15, 1401–1414. Russell, C. E., Hedger, M. P., Brauman, J. N., de Kretser, D. M., and Phillips, D. J. (1999). Activin A regulates growth and acute phase proteins in the human liver cell line, HepG2. Mol. Cell. Endocrinol. 148, 129–136. Sakaguchi, S., Sakaguchi, N., Shimizu, J., Yamazaki, S., Sakihama, T., Itoh, M., Kuniyasu, Y., Nomura, T., Toda, M., and Takahashi, T. (2001). Immunologic tolerance maintained by CD25þ CD4þ regulatory T cells: Their common role in controlling autoimmunity, tumor immunity, and transplantation tolerance. Immunol. Rev. 182, 18–32. Salogni, L., Musso, T., Bosisio, D., Mirolo, M., Jala, V. R., Haribabu, B., Locati, M., and Sozzani, S. (2009). Activin A induces dendritic cell migration through the polarized release of CXC chemokine ligands 12 and 14. Blood 113, 5848–5856. Sant, A. J., Beeson, C., McFarland, B., Cao, J., Ceman, S., Bryant, P. W., and Wu, S. (1999). Individual hydrogen bonds play a critical role in MHC class II: Peptide interactions: Implications for the dynamic aspects of class II trafficking and DM-mediated peptide exchange. Immunol. Rev. 172, 239–253. Schneider, O., Nau, R., and Michel, U. (2000). Comparative analysis of follistatin-, activin beta A- and activin beta B-mRNA steady-state levels in diverse porcine tissues by multiplex S1 nuclease analysis. Eur. J. Endocrinol. 142, 537–544. Schneyer, A., Schoen, A., Quigg, A., and Sidis, Y. (2003). Differential binding and neutralization of activins A and B by follistatin and follistatin like-3 (FSTL-3/FSRP/ FLRG). Endocrinology 144, 1671–1674. Schwall, R. H., and Lai, C. (1991). Erythroid differentiation bioassays for activin. Methods Enzymol. 198, 340–346. Schwall, R. H., Robbins, K., Jardieu, P., Chang, L., Lai, C., and Terrell, T. G. (1993). Activin induces cell death in hepatocytes in vivo and in vitro. Hepatology 18, 347–356. Scicchitano, M. S., McFarland, D. C., Tierney, L. A., Boyce, R. W., Frazier, K. S., Schwartz, L. W., and Thomas, H. C. (2008). Role of p38 in regulation of hematopoiesis: Effect of p38 inhibition on cytokine production and transcription factor activity in human bone marrow stromal cells. Blood Cells Mol. Dis. 40, 370–380.
294
Mark P. Hedger et al.
Scutera, S., Riboldi, E., Daniele, R., Elia, A. R., Fraone, T., Castagnoli, C., Giovarelli, M., Musso, T., and Sozzani, S. (2008). Production and function of activin A in human dendritic cells. Eur. Cytokine Netw. 19, 60–68. Segerer, S. E., Mu¨ller, N., Brandt, J., Kapp, M., Dietl, J., Reichardt, H. M., Rieger, L., and Ka¨mmerer, U. (2008). The glycoprotein-hormones activin A and inhibin A interfere with dendritic cell maturation. Reprod. Biol. Endocrinol. 6, 17. Sekiyama, K., Hashimoto, O., Ushiro, Y., Adachi, C., Kikusui, T., Tanemura, K., and Hasegawa, Y. (2009). Abnormalities in aggression and anxiety in transgenic mice overexpressing activin E. Biochem. Biophys. Res. Commun. 385, 319–323. Semitekolou, M., Alissafi, T., Aggelakopoulou, M., Kourepini, E., Kariyawasam, H. H., Kay, A. B., Robinson, D. S., Lloyd, C. M., Panoutsakopoulou, V., and Xanthou, G. (2009). Activin-A induces regulatory T cells that suppress T helper cell immune responses and protect from allergic airway disease. J. Exp. Med. 206, 1769–1785. Shao, L., Frigon, N. L., Jr., Sehy, D. W., Yu, A. L., Lofgren, J., Schwall, R., and Yu, J. (1992). Regulation of production of activin A in human marrow stromal cells and monocytes. Exp. Hematol. 20, 1235–1242. Shao, L. E., Frigon, N. L., Jr., Yu, A., Palyash, J., and Yu, J. (1998). Contrasting effects of inflammatory cytokines and glucocorticoids on the production of activin A in human marrow stromal cells and their implications. Cytokine 10, 227–235. Shimasaki, S., Koga, M., Esch, F., Cooksey, K., Mercado, M., Koba, A., Ueno, N., Ying, S. Y., Ling, N., and Guillemin, R. (1988). Primary structure of the human follistatin precursor and its genomic organization. Proc. Natl. Acad. Sci. USA 85, 4218–4222. Sidis, Y., Tortoriello, D. V., Holmes, W. E., Pan, Y., Keutmann, H. T., and Schneyer, A. L. (2002). Follistatin-related protein and follistatin differentially neutralize endogenous vs. exogenous activin. Endocrinology 143, 1613–1624. Smith-Garvin, J. E., Koretzky, G. A., and Jordan, M. S. (2009). T cell activation. Annu. Rev. Immunol. 27, 591–619. Sprent, J., and Schaefer, M. (1989). Antigen-presenting cells for unprimed T cells. Immunol. Today 10, 17–23. Stewart, A. G., Milborrow, H. M., Ring, J. M., Crowther, C. E., and Forage, R. G. (1986). Human inhibin genes. Genomic characterisation and sequencing. FEBS Lett. 206, 329–334. Stoitzner, P., Stossel, H., Wankell, M., Hofer, S., Heufler, C., Werner, S., and Romani, N. (2005). Langerhans cells are strongly reduced in the skin of transgenic mice overexpressing follistatin in the epidermis. Eur. J. Cell Biol. 84, 733–741. Sugama, S., Takenouchi, T., Kitani, H., Fujita, M., and Hashimoto, M. (2007). Activin as an anti-inflammatory cytokine produced by microglia. J. Neuroimmunol. 192, 31–39. Sugatani, T., Alvarez, U. M., and Hruska, K. A. (2003). Activin A stimulates IkB-a/NFkB and RANK expression for osteoclast differentiation, but not AKT survival pathway in osteoclast precursors. J. Cell. Biochem. 90, 59–67. Sugino, K., Kurosawa, N., Nakamura, T., Takio, K., Shimasaki, S., Ling, N., Titani, K., and Sugino, H. (1993). Molecular heterogeneity of follistatin, an activin-binding protein. Higher affinity of the carboxyl-terminal truncated forms for heparan sulfate proteoglycans on the ovarian granulosa cell. J. Biol. Chem. 268, 15579–15587. Sugiyama, M., Ichida, T., Sato, T., Ishikawa, T., Matsuda, Y., and Asakura, H. (1998). Expression of activin A is increased in cirrhotic and fibrotic rat livers. Gastroenterology 114, 550–558. Takahashi, S., Uchimaru, K., Harigaya, K., Asano, S., and Yamashita, T. (1992). Tumor necrosis factor and interleukin-1 induce activin A gene expression in a human bone marrow stromal cell line. Biochem. Biophys. Res. Commun. 188, 310–317.
Activin and Immunoregulation
295
Tanimoto, K., Yoshida, E., Mita, S., Nibu, Y., Murakami, K., and Fukamizu, A. (1996). Human activin bA gene. Identification of novel 50 exon, functional promoter, and enhancers. J. Biol. Chem. 271, 32760–32769. Tannetta, D. S., Muttukrishna, S., Groome, N. P., Redman, C. W., and Sargent, I. L. (2003). Endothelial cells and peripheral blood mononuclear cells are a potential source of extraplacental activin A in preeclampsia. J. Clin. Endocrinol. Metab. 88, 5995–6001. ten Dijke, P., and Hill, C. S. (2004). New insights into TGF-b-Smad signalling. Trends Biochem. Sci. 29, 265–273. Thompson, A. G., and Thomas, R. (2002). Induction of immune tolerance by dendritic cells: Implications for preventative and therapeutic immunotherapy of autoimmune disease. Immunol. Cell Biol. 80, 509–519. Thompson, D. A., Cronin, C. N., and Martin, F. (1994). Genomic cloning and sequence analyses of the bovine a-, bA- and bB-inhibin/activin genes. Identification of transcription factor AP-2-binding sites in the 50 -flanking regions by DNase I footprinting. Eur. J. Biochem. 226, 751–764. Thompson, T. B., Cook, R. W., Chapman, S. C., Jardetzky, T. S., and Woodruff, T. K. (2004). Beta A versus beta B: Is it merely a matter of expression? Mol. Cell. Endocrinol. 225, 9–17. Thompson, T. B., Lerch, T. F., Cook, R. W., Woodruff, T. K., and Jardetzky, T. S. (2005). The structure of the follistatin:activin complex reveals antagonism of both type I and type II receptor binding. Dev. Cell 9, 535–543. Tisdale, M. J. (1997). Biology of cachexia. J. Nat. Cancer Inst. 89, 1763–1773. Tone, Y., Furuuchi, K., Kojima, Y., Tykocinski, M. L., Greene, M. I., and Tone, M. (2008). Smad3 and NFAT cooperate to induce Foxp3 expression through its enhancer. Nat. Immunol. 9, 194–202. Tsuchida, K., Arai, K. Y., Kuramoto, Y., Yamakawa, N., Hasegawa, Y., and Sugino, H. (2000). Identification and characterization of a novel follistatin-like protein as a binding protein for the TGF-b family. J. Biol. Chem. 275, 40788–40796. Tsuchida, K., Nakatani, M., Uezumi, A., Murakami, T., and Cui, X. (2008). Signal transduction pathway through activin receptors as a therapeutic target of musculoskeletal diseases and cancer. Endocr. J. 55, 11–21. Turner, I. M., Saunders, P. T., Shimasaki, S., and Hillier, S. G. (1989). Regulation of inhibin subunit gene expression by FSH and estradiol in cultured rat granulosa cells. Endocrinology 125, 2790–2792. Tuuri, T., Era¨maa, M., Hilde´n, K., and Ritvos, O. (1994). The tissue distribution of activin bA- and bB-subunit and follistatin messenger ribonucleic acids suggests multiple sites of action for the activin-follistatin system during human development. J. Clin. Endocrinol. Metab. 78, 1521–1524. Tuuri, T., Era¨maa, M., Van Schaik, R. H., and Ritvos, O. (1996). Differential regulation of inhibin/activin a- and bA-subunit and follistatin mRNAs by cyclic AMP and phorbol ester in cultured human granulosa-luteal cells. Mol. Cell. Endocrinol. 121, 1–10. Uchimaru, K., Motokura, T., Takahashi, S., Sakurai, T., Asano, S., and Yamashita, T. (1995). Bone marrow stromal cells produce and respond to activin A: Interactions with basic fibroblast growth factor and platelet-derived growth factor. Exp. Hematol. 23, 613–618. Ueno, N., Ling, N., Ying, S. Y., Esch, F., Shimasaki, S., and Guillemin, R. (1987). Isolation and partial characterization of follistatin: A single-chain Mr 35,000 monomeric protein that inhibits the release of follicle-stimulating hormone. Proc. Natl. Acad. Sci. USA 84, 8282–8286. Van Pottelberge, G. R., Bracke, K. R., Demedts, I. K., De Rijck, K., Reinartz, S. M., van Drunen, C. M., Verleden, G. M., Vermassen, F. E., Joos, G. F., and Brusselle, G. G. (2010). Selective accumulation of langerhans-type dendritic cells in small airways of patients with COPD. Respir. Res. 11, 35.
296
Mark P. Hedger et al.
Vassalli, A., Matzuk, M. M., Gardner, H. A., Lee, K. F., and Jaenisch, R. (1994). Activin/ inhibin bB subunit gene disruption leads to defects in eyelid development and female reproduction. Genes Dev. 8, 414–427. Wada, W., Kuwano, H., Hasegawa, Y., and Kojima, I. (2004). The dependence of transforming growth factor-b-induced collagen production on autocrine factor activin A in hepatic stellate cells. Endocrinology 145, 2753–2759. Wakefield, L. M., Smith, D. M., Flanders, K. C., and Sporn, M. B. (1988). Latent transforming growth factor-b from human platelets. A high molecular weight complex containing precursor sequences. J. Biol. Chem. 263, 7646–7654. Wang, S. Y., Tai, G. X., Zhang, P. Y., Mu, D. P., Zhang, X. J., and Liu, Z. H. (2008). Inhibitory effect of activin A on activation of lipopolysaccharide-stimulated mouse macrophage RAW264.7 cells. Cytokine 42, 85–91. Wang, Y., Cui, X., Tai, G., Ge, J., Li, N., Chen, F., Yu, F., and Liu, Z. (2009). A critical role of activin A in maturation of mouse peritoneal macrophages in vitro and in vivo. Cell. Mol. Immunol. 6, 387–392. Wankell, M., Kaesler, S., Zhang, Y. Q., Florence, C., Werner, S., and Duan, R. (2001a). The activin binding proteins follistatin and follistatin-related protein are differentially regulated in vitro and during cutaneous wound repair. J. Endocrinol. 171, 385–395. Wankell, M., Munz, B., Hu¨bner, G., Hans, W., Wolf, E., Goppelt, A., and Werner, S. (2001b). Impaired wound healing in transgenic mice overexpressing the activin antagonist follistatin in the epidermis. EMBO J. 20, 5361–5372. Wilms, H., Schwark, T., Brandenburg, L. O., Sievers, J., Dengler, R., Deuschl, G., and Lucius, R. (2010). Regulation of activin A synthesis in microglial cells: Pathophysiological implications for bacterial meningitis. J. Neurosci. Res. 88, 16–23. Wilson, K. M., Smith, A. I., and Phillips, D. J. (2006). Stimulatory effects of lipopolysaccharide on endothelial cell activin and follistatin. Mol. Cell. Endocrinol. 253, 30–35. Winnall, W. R., Muir, J. A., and Hedger, M. P. (2009). Differential responses of epithelial Sertoli cells of the rat testis to Toll-Like receptor 2 and 4 ligands: Implications for studies of testicular inflammation using bacterial lipopolysaccharides. Innate Immun. 10.1177/ 1753425909354764. Wohlfahrt, J. G., Kunzmann, S., Menz, G., Kneist, W., Akdis, C. A., Blaser, K., and Schmidt-Weber, C. B. (2003). T cell phenotype in allergic asthma and atopic dermatitis. Int. Arch. Allergy Immunol. 131, 272–282. Woodruff, T., Krummen, L., Baly, D., Garg, S., Allison, D., Sadick, M., Wong, W., Mather, J., and Soules, M. (1993). Quantitative two-site enzyme-linked immunosorbent assays for inhibin A, activin A and activin B. Hum. Reprod. 8(Suppl. 2), 133–137. Wrana, J. L., and Attisano, L. (2000). The Smad pathway. Cytokine Growth Factor Rev. 11, 5–13. Wu, T., Patel, H., Mukai, S., Melino, C., Garg, R., Ni, X., Chang, J., and Peng, C. (2000). Activin, inhibin, and follistatin in zebrafish ovary: Expression and role in oocyte maturation. Biol. Reprod. 62, 1585–1592. Yamashita, T., Takahashi, S., and Ogata, E. (1992). Expression of activin A/erythroid differentiation factor in murine bone marrow stromal cells. Blood 79, 304–307. Yamashita, N., Nakajima, T., Takahashi, H., Kaneoka, H., Mizushima, Y., and Sakane, T. (1993). Effects of activin A on IgE synthesis and cytokine production by human peripheral mononuclear cells. Clin. Exp. Immunol. 94, 214–219. Yamashita, S., Maeshima, A., Kojima, I., and Nojima, Y. (2004). Activin A is a potent activator of renal interstitial fibroblasts. J. Am. Soc. Nephrol. 15, 91–101. Yang, X., Letterio, J. J., Lechleider, R. J., Chen, L., Hayman, R., Gu, H., Roberts, A. B., and Deng, C. (1999). Targeted disruption of SMAD3 results in impaired mucosal immunity and diminished T cell responsiveness to TGF-b. EMBO J. 18, 1280–1291.
Activin and Immunoregulation
297
Yasuda, H., Mine, T., Shibata, H., Eto, Y., Hasegawa, Y., Takeuchi, T., Asano, S., and Kojima, I. (1993). Activin A: An autocrine inhibitor of initiation of DNA synthesis in rat hepatocytes. J. Clin. Invest. 92, 1491–1496. Yndestad, A., Ueland, T., ie, E., Florholmen, G., Halvorsen, B., Attramadal, H., Simonsen, S., Fr land, S. S., Gullestad, L., Christensen, G., Dama˚s, J. K., and Aukrust, P. (2004). Elevated levels of activin A in heart failure: Potential role in myocardial remodeling. Circulation 109, 1379–1385. Yndestad, A., Larsen, K. O., ie, E., Ueland, T., Smith, C., Halvorsen, B., Sjaastad, I., Skj nsberg, O. H., Pedersen, T. M., Anfinsen, O. G., Dama˚s, J. K., Christensen, G., et al. (2009). Elevated levels of activin A in clinical and experimental pulmonary hypertension. J. Appl. Physiol. 106, 1356–1364. Yoshida, E., Tanimoto, K., Murakami, K., and Fukamizu, A. (1998). Isolation and characterization of 50 -regulatory region of mouse activin bA subunit gene. Biochem. Mol. Biol. Int. 44, 325–332. Yu, J., Shao, L., Vaughan, J., Vale, W., and Yu, A. L. (1989). Characterization of the potentiation effect of activin on human erythroid colony formation in vitro. Blood 73, 952–960. Yu, A. W., Shao, L. E., Frigon, N. L., Jr., and Yu, J. (1994). Detection of functional and dimeric activin A in human marrow microenvironment. Implications for the modulation of erythropoiesis. Ann. NY Acad. Sci. 718, 285–298. (discussion 298–289). Yu, J., Shao, L. E., Frigon, N. L., Jr., Lofgren, J., and Schwall, R. (1996). Induced expression of the new cytokine, activin A, in human monocytes: Inhibition by glucocorticoids and retinoic acid. Immunology 88, 368–374. Yu, E. W., Dolter, K. E., Shao, L. E., and Yu, J. (1998). Suppression of IL-6 biological activities by activin A and implications for inflammatory arthropathies. Clin. Exp. Immunol. 112, 126–132. Zhang, L., Deng, M., Parthasarathy, R., Wang, L., Mongan, M., Molkentin, J. D., Zheng, Y., and Xia, Y. (2005a). MEKK1 transduces activin signals in keratinocytes to induce actin stress fiber formation and migration. Mol. Cell. Biol. 25, 60–65. Zhang, X. J., Li, Y., Tai, G. X., Xu, G. Y., Zhang, P. Y., Yang, Y., Lao, F. X., and Liu, Z. H. (2005b). Effects of activin A on the activities of the mouse peritoneal macrophages. Cell. Mol. Immunol. 2, 63–67. Zhou, J., Tai, G., Liu, H., Ge, J., Feng, Y., Chen, F., Yu, F., and Liu, Z. (2009). Activin A down-regulates the phagocytosis of lipopolysaccharide-activated mouse peritoneal macrophages in vitro and in vivo. Cell. Immunol. 255, 69–75. Zimmers, T. A., Davies, M. V., Koniaris, L. G., Haynes, P., Esquela, A. F., Tomkinson, K. N., McPherron, A. C., Wolfman, N. M., and Lee, S. J. (2002). Induction of cachexia in mice by systemically administered myostatin. Science 296, 1486–1488. Zipori, D., and Barda-Saad, M. (2001). Role of activin A in negative regulation of normal and tumor B lymphocytes. J. Leukoc. Biol. 69, 867–873.
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Feedback Regulation by Inhibins A and B of the Pituitary Secretion of Follicle-Stimulating Hormone Yogeshwar Makanji, Craig A. Harrison, and David M. Robertson Contents I. Introduction II. Structure, Mechanism of Action, and Function of Inhibins and Activins A. Structure B. Mechanism of action C. Function III. Inhibins A and B Mediated FSH Suppression in Males and Females A. Human menstrual cycle B. Relationship between inhibin B and FSH C. Relationship between inhibin A and FSH D. Relationship between AMH, inhibins, and FSH E. Relationship between inhibins and LH F. Inhibins and regulation of FSH in the male IV. Understanding the Increased Potency of Inhibin B In Vivo and In Vitro A. The effect of glycosylation on inhibins A and B bioactivity B. The key role of betaglycan in mediating inhibins A and B biological activities C. Updated model of inhibin antagonism of activin action D. Other factors potentially contributing to the increased potency of inhibin B E. Future directions Acknowledgments References
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Abstract Inhibins A and B are gonadal factors that negatively regulate FSH synthesis by the anterior pituitary. Across the menstrual cycle, women show a strong inverse correlation between circulating FSH and inhibin B, estradiol, and antiMullerian hormone (AMH), but not with inhibin A. Estradiol is believed to provide a tonic inhibitory effect while the inhibitory role of AMH is unknown. In human males, inhibin B is the primary testicular factor regulating FSH with limited effects by gonadal steroids. In vitro and in vivo studies in rats indicate that inhibin B is more biologically active than inhibin A but showed a lower affinity for the activin type II receptors and the co-receptor, betaglycan, suggesting an alternative mechanism. While this review reinforces the important role inhibin plays in regulating FSH, the observed differences in mode of action of inhibins A and B and their interplay with other gonadal factors are still poorly understood. ß 2011 Elsevier Inc.
I. Introduction The ovarian feedback mechanism in the regulation of gonadotrophins is a complex process with both steroidal and nonsteroidal factors involved. Gonadal steroids have a profound suppressive effect on both FSH and LH but require pharmacological doses to independently suppress these pituitary hormones and, in particular, FSH into the physiological range. The original work of McCullagh (1932) and Setchell and Jacks (1974) highlighted the need for additional secreted ovarian factors in the regulation of FSH. For reviews dealing with the early studies on the respective roles of steroidal and protein factors which regulate the pituitary gonadotrophins, the reader is referred to the following references: de Kretser et al. (2002) and Woodruff and Mather (1995). Inhibins are primarily products of the ovary and testis and their recognized role to date is to inhibit FSH synthesis and secretion by the pituitary. Other roles have been reported but are less clearly defined (for recent reviews, see Itman et al., 2006; Knight and Glister, 2006). Their role is inhibitory to suppress the stimulatory action of the structurally related activins (A, AB, B). Over the past 20 years and with the development of specific immunoassays for serum inhibins A and B, an understanding of the feedback regulation of FSH in vivo and to a lesser extent LH has now been developed. This review will present recent data on the modes of action of inhibins A and B and explore their relationships with pituitary gonadotrophins in various physiological reproductive states.
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II. Structure, Mechanism of Action, and Function of Inhibins and Activins A. Structure Inhibins are produced as heterodimers of b-subunits with the common inhibin a-subunit to form inhibin A (a-bA) and inhibin B (a-bB). The subunits dimerize via a single cysteine linkage, and the precursor form (Fig. 14.1A) undergoes a series of proteolytic cleavages by the serine peptidase, furin, at the C-terminal region, to give rise to a heterodimer A
* Pro
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a233STPLMSWPWSPSALRLLQRPPEEPAAHANCHRVALNISFQELGWERWIVYPPSFIFHYCHGGCGLHIPP bA -----------------311GLECDGKVNICCKKQFFVSFKDIGWNDWIIAPSGYHANYCEGECPSHIAG bB -----------------293GLECDGRTNLCCRQQFFIDFRLIGWNDWIIAPTGYYGNYCEGSCPAYLAG a
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bA TSGSSLSFHSTVINHYRMRGHSPFANLKSCC—-VPTKLRPMSMLYYDDGQNIIKKDIQNMIVEE-CGCS bB VPGSASSFHTAVVNQYRMRGLNP-GTVNSCC—-IPTKLSTMSMLYFDDEYNIVKRDVPNMIVEE-CGCA
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Figure 14.1 (A) Schematic diagram of the precursor inhibin molecule. The cleavage sites are denoted by (^) and (*) represents known glycosylation sites. (B) Amino acid sequence alignments (aligned based on conserved cysteines) of the C-terminal mature domains of the inhibin a-, bA-, and bB-subunits. bA- and bB-subunits show 63% homology; nonconserved residues are in bold. (B) A homology model of the mature 34-kDa inhibin molecule based on the 3D structure of activin A and BMP-3 and BMP-6 structures (Allendorph et al., 2007; Harrington et al., 2006; Thompson et al., 2003). Both inhibins A and B are believed to show a similar 3D structure. The a-subunits are glycosylated preferentially at two sites marked Asn268 and Asn302.
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with a molecular mass of 31–34 kDa (Antenos et al., 2007; Makanji et al., 2007). Homodimers of the b-subunits also exist as activins A, B, or AB (Bilezikjian et al., 2006; Thompson et al., 2004). Since inhibins A and B share a common a-subunit, their differences lie in their b-subunits with 63% homology between the bA- and bB-subunits (Fig. 14.1B). The three-dimensional structures of inhibins A or B have yet to be determined. Therefore, we recently constructed a homology model of the inhibin a-subunit (Fig. 14.1C) based on the activin A and BMP-3 and BMP-6 structures (Allendorph et al., 2007; Harrington et al., 2006; Thompson et al., 2003). Seven cysteine residues can be aligned between these TGFb family members and are thought to confer the same overall conformation in the a-subunit as observed in the activin and BMP structures. Thus within the a-subunit, two pairs of anti-parallel b-strands, forming first a short and then a long “finger” stretch outward from the cysteine knot core of the monomer. The characteristic curvature of these fingers creates concave and convex surfaces through this region. At the opposite end of the molecule, a high proline content (8 of 24 residues) would ensure the inhibin a-subunit has a truncated a-helix relative to other TGFb family members (Fig. 14.1C). Unlike other species, the human inhibin a-subunit has two N-linked glycosylation sites; at Asn268 and Asn302, which are primarily responsible for the molecular mass heterogeneity observed with human inhibins A and B forms. Asn268 is always glycosylated (monoglycosylated 31-kDa inhibin A or B), whereas Asn302 is differentially glycosylated (diglycosylated 34-kDa inhibin A or B) (Makanji et al., 2007; Mason et al., 1996). Glycosylation of Asn268 is required for synthesis of inhibin (Antenos et al., 2007) and potentially for its high-affinity binding to betaglycanþActRIIA/B, whereas glycosylation of Asn302 results in a reduced affinity for receptors and reduced biological activity of both inhibins A and B (Makanji et al., 2007). The physiological impact of these glycosylation events will be discussed in later sections. Based on sequence homology, inhibins belong to the TGFb superfamily of growth and differentiation factors. Ligands within this family can be divided into the TGFb, bone morphogenetic protein (BMP), and activin/ inhibin subgroups. Sequence identity is 30–50% for ligands in different subgroups and 60–85% for ligands in the same subgroup. The three TGFb isoforms, TGFb1, 2, and 3, have identities between 74% and 82% and yet are thought to play distinct physiological roles in a variety of target tissues (Thompson et al., 2004). Isoform-specific actions can be partially attributed to differential expression patterns, but evidence is mounting that minor differences in signaling mechanisms and binding affinities of each TGFb isoform may influence the degree and kind of signal transduction that is engaged in a target cell (Thompson et al., 2004). Therefore, sequence variation within a ligand subgroup may be more important in determining
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the biological activity of specific TGFb superfamily ligands than originally surmised. Our recent structure/function analysis of inhibins A and B (62% amino acid identity between b-subunits) supports this concept.
B. Mechanism of action 1. Activins A and B signal transduction It is recognized that inhibins regulate FSH secretion by inhibiting the stimulatory actions of the structurally related proteins, activins (Bilezikjian et al., 2004). Activin signal transduction is mediated by binding to transmembrane serine/threonine kinase receptors. Activation of these receptors results in the transmission of downstream gene modulation which is mediated by Smad transcription factors (Massague et al., 2005). To date four activin receptors have been identified, activin type IB (ALK 4), type IC (ALK 7), type IIA, and type IIB (ActRIIA/B) (see reviews Harrison et al., 2005; Shi and Massague, 2003). Like TGFb, activins adopt a cooperative mode of ligand binding. Activin type II receptors can bind ligand independently of the type I receptor; however, they are unable to mediate signaling (De Winter et al., 1996; Harrison et al., 2004). Type I receptors for activin can only recognize ligand that is bound to the type II receptor. Consequently, the ligand–type II receptor recruits and phosphorylates the type I receptor and mediates downstream signaling via Smad 2, 3, and 4 (Harrison et al., 2005). Recent studies have shown that activin B signals via ALK7 and ALK4 whilst activin A preferentially mediates its signaling via ALK4 and not ALK7 (Bernard et al., 2006; Tsuchida et al., 2004). ALK7 is expressed in both the rat pituitary and the mouse gonadotrope (LbT2) cell line; tissue/ cell systems currently used to understand inhibins A and B biological activity. Thus, the inhibitory roles of inhibins A and B may depend on the presence of the appropriate activin form and receptor subtypes in the target tissues, like the pituitary. 2. Current model of inhibin antagonism of activin action Inhibin’s mechanism of action is attributed to the functional antagonism of activin signaling. The key components of this antagonism are activin type II receptors and most importantly the coreceptor, betaglycan. It has been shown that betaglycan is required for the formation of a high-affinity complex consisting of inhibin A, ActRIIA/B, and betaglycan (Lewis et al., 2000; Makanji et al., 2008; Wiater et al., 2009). Gonadotropes are the primary site of action for inhibin and activin in the pituitary and these cells have been shown to express the mRNA and protein for the inhibin coreceptor, betaglycan (Chapman and Woodruff, 2003; MacConell et al., 2002), ActRIIA/B, ALK4 (Dalkin et al., 1996), and ALK7 (Bernard et al., 2006). HEK293 cells transiently transfected with ActRII alone had a low
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affinity (K ¼ 6.3 nM) for inhibin A but this affinity was increased 30-fold (K ¼ 0.2 nM) when ActRII and betaglycan were cotransfected (Lewis et al., 2000). The role of betaglycan in inhibin B’s mode of action will be discussed in later sections.
C. Function 1. Regulation of FSHb subunit synthesis by activins and inhibins Activin bB subunit synthesis is under the transcriptional control of GnRH and gonadal steroids (Melamed, 2010; Thackray et al., 2010; Xia and Schneyer, 2009). In vitro studies using rat pituitary cell cultures have shown that activin stimulates (Attardi and Miklos, 1990) and inhibin (Attardi et al., 1992) suppresses the levels of FSHb subunit mRNA and FSH protein (Carroll et al., 1989). In addition, inhibin (Attardi and Winters, 1993) decreases and activin increases the half-life of FSHb subunit mRNA (Carroll et al., 1991). Activin directly stimulates the synthesis of the FSHb subunit gene (Ling et al., 1986; Weiss et al., 1995) via Smad 2, 3 downstream signaling (Bernard, 2004). It was shown that gonadal factors and not GnRH suppress ActRIIA receptor expression in the rat pituitary as evidenced by the increase in FSHb subunit mRNA and FSH protein postovariectomy (Dalkin et al., 1994). Thus, gonadal inhibins may also downregulate the expression of the ActRIIA receptor, thereby suppressing activin-mediated FSH secretion. In rodents, the steroids (progesterone and corticosterone) stimulate the secondary FSH surge during the rat estrous cycle by preferentially stimulating activin B production compared to inhibins/follistatin (Tebar et al., 2000). It has been proposed that activin B and not activin A is the major pituitary form stimulating FSH and perhaps LH synthesis and secretion in rodents and humans. Several studies utilizing rat pituitary cell cultures (Bilezikjian et al., 1993; Corrigan et al., 1991) and mouse cell lines (LbT2) (Pernasetti et al., 2001) suggest that activin B is the main form stimulating FSHb subunit mRNA and protein expression. In support, activin bB-subunit is detected in human pituitaries, while activin bΑ-subunit is not (Uccella et al., 2000). Thus, as observed in mice and rats, activin B may also be the predominant activin form stimulating FSH release from the human anterior pituitary. Identifying the active activin form in human pituitaries is important when activin B can signal via ALK4 and ALK7 (Bernard et al., 2006), whilst activin A can only signal via ALK4 (Tsuchida et al., 2004). However, it has been proposed that activin B preferentially mediates its pituitary actions via ALK4 compared to ALK7 as it may have a lower affinity for ALK7 (Bernard et al., 2006). Thus, antagonism of activin B action at the pituitary by inhibins A and B must take into account the specificity and affinities of the activin receptor subtypes present.
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2. Regulation of LHb subunit synthesis by activins and inhibins Recent studies suggest that activins also play a role in the regulation of the LHb subunit. In vitro studies in the mouse gonadotrope cell line, LbT2, showed that activin A was able to increase LHb subunit mRNA and LH levels (Yamada et al., 2004). In combination with GnRH, activin A is a potent activator of the LHb subunit promoter in LbT2 cells (Coss et al., 2005; Yamada et al., 2004). In Smad 3 deficient mice, both LHb and FSHb subunit levels are reduced attributed to impaired activin signaling, while overexpression of Smad 7 (inhibitor of activin signaling) in LbT2 cells resulted in abolishment of activin-induced LHb subunit expression (Coss et al., 2005). The role of inhibin in regulating LH is unclear.
III. Inhibins A and B Mediated FSH Suppression in Males and Females The development of specific ELISAs for measuring inhibins A and B in blood (Groome et al., 1996) has enabled studies to be undertaken to explore the roles of circulating inhibins in the regulation of pituitary gonadotrophins in humans. In the sections below, data from the past 10 years will be emphasized.
A. Human menstrual cycle As seen in Fig. 14.2, serum inhibins A and B show independent patterns over the menstrual cycle. Under FSH stimulation (Groome et al., 1996; Welt et al., 2001), inhibin B produced by granulosa cells of small antral follicles increases early in the follicular phase and then dips toward midcycle. As part of the proposed feedback mechanism, the decline in inhibin B is attributed in turn to its inhibitory effects on pituitary FSH synthesis (Welt et al., 2001). In the luteal phase, inhibin B levels are lower, most likely attributed to the limited production by the early follicle stages of the next follicular wave. Inhibin A, however, is primarily a product of granulosa cells from antral follicles and luteal cells of the corpus luteum and shows a progressive increase in the follicular phase in parallel with the development of the dominant follicle. The pattern of inhibin A parallels that of estradiol, a known FSH-sensitive follicle product. Exogenous FSH stimulation in the follicular phase of women leads to increases in both inhibins A and B (Burger, 1992; Burger et al., 1998; Eldar-Geva et al., 2000; Welt and Schneyer, 2001) supporting the positive arm of this feedback mechanism. Overall, these data support a reciprocal relationship between inhibin B and FSH in the follicular phase as supported by several investigators (Welt and Schneyer, 2001; Welt et al., 2003), but a less clear relationship is evident
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Figure 14.2 The changing hormone patterns of ovarian and pituitary hormones in women with ovulatory menstrual cycles during mid-reproductive age (MRA; 25–35 years) and in women approaching menopause (aged 45–55 years) (Robertson et al., 2008). A classification system (Cycle Type 1–3) was used to describe changes in circulating hormone patterns in women in the late reproductive aged group.
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between inhibin A and FSH. In an attempt to clarify in more detail the relationships in serum ovarian and pituitary hormones across the menstrual cycle, a large study was undertaken in women during their reproductive life (21–35 years) and approaching menopause (45–55 years) (Hale et al., 2007). The rationale for including women late in their reproductive life was to explore the changing relationships between ovarian and pituitary hormones as the ovary is deleted of follicles with a corresponding decrease in ovarian feedback factors on the pituitary. The following observations were made (Robertson et al., 2008, 2009): (a) Ovulatory cycles with similar patterns of FSH, LH, inhibins A and B, estradiol, and progesterone compared to women during their midreproductive life were observed late into reproductive life. Serum AMH decreased in parallel with the known decrease in ovarian follicle number with age (Fig. 14.2). (b) The first observed changes approaching menopause were decreases in serum inhibin B and AMH in the follicular phase and a concomitant increase in serum FSH without changes in serum LH, inhibin A, estradiol, and progesterone in either phase of the cycle (Fig. 14.2). The fall in both inhibin B and AMH is believed to reflect a decline in ovarian follicle number and the rise in FSH is attributed to the reduction in ovarian feedback on the pituitary.
B. Relationship between inhibin B and FSH To clarify which of these ovarian hormones is independently correlated with FSH, a multiple linear regression analysis was undertaken whereby comparisons were undertaken within the early, mid, and late phases of the follicular phase across both the young and older aged women. Surprisingly, serum estradiol, AMH and to a limited degree inhibin B, but not inhibin A inversely correlated with FSH. However, it was hypothesized that a direct comparison between these hormones within any one phase of the cycle was not necessarily appropriate as the time course of ovarian action on the pituitary may span several days. The data were then reanalyzed whereby multiple linear regression analyses were undertaken between serum ovarian hormone levels and serum FSH (and LH) 3 days later (lagged analyses) Cycle Type 1 shows similar hormone profiles (other than AMH) to that seen in women in their mid-reproductive years (MRA); Cycle Type 2 shows cycles in women in which serum inhibin B decreases and FSH increases while inhibin A, estradiol LH remain unchanged. These are believed to be the first hormonal changes seen approaching menopause. Cycle Type 3 shows cycles in women with ovulatory cycles where additional hormonal changes are observed leading to distorted profiles. Copyright to the North American Menopause Society.
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within both follicular and luteal phases. The results obtained showed that serum inhibin B and FSH were highly correlated and so were serum AMH and FSH, but not serum estradiol or inhibin A with FSH (Fig. 14.3). FSH Follicular phase Luteal phase Estradiol Inhibin A Inhibin B Progesterone AMH (Lagged)
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Figure 14.3 Slope values and 95% confidence limits for regression lines between serum ovarian hormones and plasma FSH and LH in women over the ovulatory menstrual cycle (as presented in Fig. 14.2). The lagged data refer to analyses whereby the ovarian hormones are compared with serum FSH and LH 3 days later. Slope values greater or less than 1 refer to a direct or inverse relationship; see text for further details (Robertson et al., 2009). Copyright to the Endocrine Society.
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The inverse relationship observed between estradiol and FSH between regions of the menstrual cycle but not across regions within cycles suggests that estradiol sets an overall inhibitory tone in regulating FSH levels across the cycle but does not play a role as an independent factor in regulating FSH within the cycle. However, estradiol showed a direct relationship with LH within cycle in the follicular phase. Other studies in which women were treated with the estrogen antagonist, tamoxifen concluded that the inhibitory effect of estradiol on FSH was primarily during the luteal phase and luteal-follicular transition rather than during the follicular phase of the menstrual cycle (Welt et al., 2003).
C. Relationship between inhibin A and FSH It was surprising that there was such a poor relationship between serum inhibin A and FSH in both follicular and to a lesser extent, luteal phases. One possibility was that the serum levels of inhibin A relative to inhibin B were much lower across the menstrual cycle and thus failed to show a significant association in vivo. In our recent study using inhibins A and B reference preparations calibrated in terms of their amino acid content (Makanji et al., 2009), the serum levels of inhibins A and B levels were noted in fact to be similar across the follicular phase. We concluded that in humans, inhibin A is considerably less active in suppressing FSH than inhibin B similar to conclusions derived from in vivo studies in the human (Welt et al., 2003) and macaque (Fraser et al., 1999) based on the effects of estrogen antagonists on circulating reproductive hormone levels. Studies in prepubertal girls showed a parallel increase with age in serum inhibin A, inhibin B, and FSH reaching a plateau between 14 and 18 years (Crofton et al., 2002a). These increases reflect the stimulatory effects of FSH on follicular development and inhibins A and B synthesis and secretion increasing to a stage when the ovarian feedback mechanisms are able to effectively regulate FSH.
D. Relationship between AMH, inhibins, and FSH The highly significant inverse relationship between serum AMH and FSH (but not LH) in either lagged or nonlagged analyses was a surprise. Studies in the mouse gonadotrope (LbT2) cell line showed AMH stimulated an increase, not a decrease in FSH (Bedecarrats et al., 2003). Similarly, a limited increase in FSH was observed in AMH transgenic mice while AMH null mice showed a limited decrease (reviewed in Bedecarrats et al., 2003; Visser et al., 2006). These findings suggest that AMH is positively correlated with FSH at least in mice. In contrast, there is evidence that AMH has an inhibitory role on FSH action in the ovary. AMH inhibits FSH-induced ovarian follicle growth in vitro as well as inhibiting FSH-stimulated
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expression of aromatase (di Clemente et al., 1994; Visser et al., 2006). In AMH null mice, age-related depletion of follicles is accelerated which is attributed to a rapid recruitment of early stage follicles despite lower serum FSH levels (Durlinger et al., 1999; Visser et al., 2006). Overall, these studies support the notion that AMH has an inhibitory role on FSH action on ovarian folliculogenesis and that with age and the loss of follicles, the inhibitory role of AMH on the ovary is lifted. How thus to explain the inverse independent association between serum AMH and FSH in the human studies discussed above? There are several hypotheses: (a) AMH is a surrogate for the number of developing follicles and that a similar pattern would exist if AMH concentration was replaced with follicle number in the above analyses. (b) AMH potentiates FSH-stimulated inhibin bB subunit synthesis by the ovary as seen with other members of the TGFb family (BMP2 ( Jaatinen et al., 2002), TGFb1/2, GDF9 (Shi et al., 2009)) which act through the ALK2/3/6 receptor pathway. Thus inhibin B production by the ovary would be directly stimulated by AMH. When AMH levels decrease with age due to a reduction in follicle number, inhibin B levels decline also; and as a consequence, FSH rises. Thus AMH is indirectly inversely associated with FSH through its stimulatory action on inhibin B synthesis by the ovary. In support, while there is a significant direct relationship between AMH and inhibin B in the follicular phase (r ¼ 0.60, p < 0.001) (Robertson et al., 2009), there is no such relationship between AMH and inhibin A (r ¼ 0.11 ns EF/MF) as inhibin A is stimulated through an alternative (PKA) pathway (Hua et al., 2008). What is unclear is to what extent AMH potentiates inhibin B’s action if this hypothesis is correct? And to extend this argument, are similar effects on inhibin B and thus FSH seen with other members of the TGFb family (BMP2, TGFb1/2, GDF9)? (c) AMH has a direct and independent effect on pituitary FSH but not LH by an unknown mechanism.
E. Relationship between inhibins and LH One interesting finding (Robertson et al., 2009) was that inhibin B was independently inversely correlated with LH in the follicular phase. In fact, of all the parameters analyzed, inhibin B was the only factor which inversely correlated with LH while inhibin A and estradiol (which were directly correlated) and AMH showed no significant correlations at all. Progesterone did show an inhibitory relationship in the follicular phase but its contributions are likely to be minor. In contrast, the inhibitory effect of progesterone on LH in the luteal phase is considerable. These data provide in vivo support of in vitro
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studies discussed in sections above where inhibins inhibit LH synthesis and secretion. To what extent inhibin B affects the biological roles of LH across the menstrual cycle is unclear but as observed in Fig. 14.2, despite unchanged serum estradiol levels throughout then menstrual cycle with ageing, serum LH levels increase. This increase may also reflect a change in hypothalamic input rendering the pituitary more sensitive to GnRH (Sealfon et al., 1990) but a direct inverse relationship by inhibins must also be considered.
F. Inhibins and regulation of FSH in the male Similar to that seen in the female, it is generally agreed that the testicular regulation of FSH and LH involves both gonadal steroids (testosterone/ estradiol) and inhibin B (Hayes et al., 2001; Plant and Marshall, 2001). Inhibin A levels are not detectable by ELISA in the adult male. Contraceptive levels of testosterone and progesterone analogues can suppress serum FSH by 95% and LH by >98% (McLachlan et al., 2002). However, based on studies using the steroid synthesis antagonist, ketoconazole in human subjects (Pitteloud et al., 2008), FSH levels increase but not to the levels comparable to that seen following castration or hemicastration. As a consequence, these results suggest that inhibin B is probably the main regulator. The inhibitory effects of exogenous testosterone and estradiol on circulating FSH were investigated in ketoconazole treated-men (Pitteloud et al., 2008). Estradiol but not testosterone was able to suppress FSH in support of earlier studies (Hayes et al., 2001). Similar findings were obtained in men with GnRH insufficiency confirming the direct inhibitory role of estradiol on pituitary FSH secretion (Pitteloud et al., 2008). The stimulation of Sertoli cell inhibin by FSH both in vivo and in vitro appears to be complex. FSH stimulates inhibin a-subunit and to a lesser extent, bB-subunit in Sertoli cells in vitro with a limited increase (Anderson, 2001) following exogenous administration in circulating inhibin B in vivo in humans. A 95% reduction in serum FSH in fertile men by contraceptive steroid doses lead to a 70% suppression in inhibin B synthesis. However, inhibin B levels are markedly suppressed in men with primary testicular failure or following irradiation or chemotherapy implicating the role of earlier germ cells in the production of inhibin B (Anderson, 2001). It is hypothesized that a germ cell factor in conjunction with FSH is required to promote inhibin B synthesis by the Sertoli cell (Anderson and Sharpe, 2000). In Sertoli cell cocultures with germ cells, pachytene spermatocytes suppressed FSH-induced bB-subunit mRNA and inhibin B levels without affecting inhibin a-subunit mRNA (Clifton et al., 2002); however, the identity of this inhibitory factor is not known. In a similar developmental pattern to that seen in girls, inhibin B levels showed a positive correlation with FSH during prepuberty and early puberty in boys, while in late puberty through to adulthood, inhibin B is
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inversely correlated with FSH (Andersson and Skakkebaek, 2001; Crofton et al., 1997, 2002b).
IV. Understanding the Increased Potency of Inhibin B In Vivo and In Vitro Analysis of the interactions between the ovarian inhibins and pituitary FSH and LH across the menstrual cycle and with age raised questions about the respective roles of inhibins A and B in this process. It was surprising to note that based on regression analyses, inhibin A, in contrast to inhibin B, appeared to play a minor role in regulating FSH across the menstrual cycle. Below, we summarize our most recent studies to provide a basis why inhibin B is more bioactive than inhibin A in vivo and in vitro.
A. The effect of glycosylation on inhibins A and B bioactivity Recently, we showed that highly purified preparations of recombinant human 31-kDa inhibin B were more bioactive (4.2- and 1.5-fold) than 31-kDa inhibin A at suppressing FSH, in vitro and in vivo, respectively (Fig. 14.4A) (Makanji et al., 2009). We attributed the less marked in vivo difference (1.5-fold increase) to a 1.8-fold greater rate of clearance of 31kDa inhibin B compared to 31-kDa inhibin A. In addition, the monoglycosylated 31-kDa inhibins A and B forms were 2.5-fold more potent than the corresponding diglycosylated forms in vitro (Fig. 14.4A). Despite this increased bioactivity, 31- and 34-kDa inhibin B have markedly reduced affinities for their receptors (betaglycan and ActRIIA/B) compared to inhibin A. (Fig. 14.4B–D), with the diglycosylated forms showing reduced affinities compared to the mono-glycosylated forms. The presence of the second glycosylation site (Asn302) on the inhibin a-subunit is also likely to influence its in vivo bioactivity as the circulating half-life of both inhibins is prolonged (1.8-fold) presumably increasing its bioavailability. Overall, inhibin B isoforms (31- and 34-kDa) are more bioactive than the inhibin A isoforms, despite reduced affinities to ActRIIA/B and betaglycan. However, glycosylation of Asn302 results in a reduction in bioactivity probably due to diminished affinities to betaglycan and type II receptor complex. These results suggest that the mechanism of inhibin B action differs from that of inhibin A.
B. The key role of betaglycan in mediating inhibins A and B biological activities The reduced affinity of diglycosylated (34-kDa) inhibins A and B (10- to 19-fold) for betaglycan compared to the 31-kDa isoforms could be attributed to the influence of glycosylation at Asn302. The reduced affinity of
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Figure 14.4 In vitro bioactivities (A) of inhibins A and B isoforms and their relative binding activities to betaglycan (B) and betaglycanþactivin type IIA/B receptors (C and D, respectively). Note that while inhibin B is more biologically active than inhibin A, its binding to betaglycanþactivin type II receptor is markedly reduced. We suspect that an additional binding protein is needed to facilitate the biological action of inhibin B. The letters denote significant differences, p < 0.05.
31-kDa inhibin B (10-fold) for betaglycan relative to 31-kDa inhibin A was more difficult to explain and led to our characterization of the betaglycan binding site on inhibins A and B. Initially, we proposed that the b-subunits must contribute to inhibin’s affinity for betaglycan. To test this, we generated a-bA/bB subunit chimeras, whereby nonconserved bA-subunit residues (Fig. 14.1B) were incorporated into the bB-subunit. Interestingly, none of the chimeras tested had increased affinity for betaglycan relative to wild-type inhibin B (unpublished observations), suggesting that the b-subunits are not directly involved in binding to betaglycan. Subsequently, we identified an epitope for high-affinity betaglycan binding spanning the outer convex surface of the inhibin a-subunit (Makanji et al., 2008). Homology modeling indicated that key a-subunit residues (Tyr282, Val340, Thr343, Ser344, Phe350, Lys351, and Tyr352) form a contiguous epitope in this region of the molecule. Disruption of betaglycan binding by the simultaneous substitution of Thr343, Ser344, and Tyr352
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(TSY) (Fig. 14.5) to alanine yielded inhibins A and B variants that were severely compromised in their ability to suppress activin-induced FSH release by rat pituitary cells in culture. These results indicate that betaglycan binding is essential for both inhibins A and B biological activity. Since inhibins A and B both require interactions with betaglycan and the increased in vitro bioactivity of inhibin B cannot be explained by affinity for the betaglycan/ActRII complex, we hypothesize that other mechanisms (e.g., additional binding proteins) may be involved in mediating inhibin B’s biological activity. In cross-linking studies, an inhibin B-specific binding protein has been identified in the LbT2 pituitary gonadotrope cell line, which may be the inhibin B specific accessory protein, Fig. 14.6A (Makanji et al., 2009). Similar to observations in rat pituitary cell cultures, inhibin B was more bioactive than inhibin A at suppressing endogenous FSH in LbT2 cells, Fig. 14.6B (Makanji et al., 2009).
C. Updated model of inhibin antagonism of activin action Based on our recent studies, we propose the following models for inhibins A and B antagonism of activin actions: For inhibin A, betaglycan binds with high affinity to the outer convex surface of the “fingers” of the a-subunit (Makanji et al., 2008), which leads to an increase in the affinity of the bA-subunit for ActRIIA/B. (Lewis et al., Betaglycan-binding site ActRIIA/Bbinding site Tyr352 Ser344 Thr343
bA/B-subunit
Asn302 Asn268
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Figure 14.5 Schematic diagram of the binding of betaglycan and activin type II receptor to the finger regions of the a- and b-subunits of inhibin. The key residues (Thr343, Ser344, and Tyr352) involved in high-affinity betaglycan binding and inhibins A and B bioactivity are indicated by arrows on the inhibin a-subunit. The location of the glycosylation site at Asn302 of the inhibin a-subunit interferes with the binding of inhibin to the activin type II receptor, providing an explanation as to why the diglycosylated forms of inhibins A and B are less active than the monoglycosylated forms. By deduction, the putative inhibin B-binding protein is believed to the exposed residues of either inhibin a- or b-subunits.
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Figure 14.6 (A) Identification of an inhibin B-specific accessory protein. LbT2 cells were incubated with 125I-inhibin A or 125I-inhibin B for 4 h at room temperature and cross-linked using the hetero-bifunctional cross-linker BS3. Inhibin cross-linked complexes were separated by SDS-PAGE and visualized by autoradiography (Makanji et al., 2009). (B) In vitro bioactivities of 31/34-kDa inhibins A and B. Inhibin dose–response curves based on FSH suppression were determined in mouse pituitary gonadotrope (LbT2) cell line cultures (n ¼ 2). 31/34-kDa inhibin A was used as reference preparation (Makanji et al., 2009). Copyright to the Endocrine Society.
2000; Makanji et al., 2008) (Fig. 14.5). There are two possible mechanisms that could explain inhibin A’s enhanced affinity for ActRIIA/B in the presence of betaglycan: (i) an initial constraint of inhibin A at the membrane surface by its high affinity binding receptor (betaglycan) would lead to a
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decrease in the entropy and, hence, an increase in affinity for the second receptor binding event (ActRII/IIB) and (ii) inhibin A may facilitate direct interactions between the two receptor molecules. An analysis of structural data on other TGFb ligands and their receptor complexes supports a model of cooperative oligomeric receptor assembly over direct receptor–receptor interactions (Greenwald et al., 2004). Whatever the mechanism, the formation of the inhibin A/betaglycan/ActRII complex precludes activin from accessing its signaling receptors. For inhibin B, interactions with betaglycan and ActRII occur as above, and betaglycan is required to mediate inhibin B’s antagonism of activin. However, inhibin B’s affinity for the betaglycan/ActRII complex is not sufficient to explain its potency in suppressing FSH secretion by the anterior pituitary. We have observed an additional inhibin B binding protein in LbT2 gonadotrope cells that is absent in cell lines that are less responsive to inhibin B than inhibin A (unpublished observations). We propose that this pituitary protein binds inhibin B and increases its affinity for the betaglycan/ ActRII complex. In this way, inhibin B activity at the pituitary can be enhanced relative to inhibin A.
D. Other factors potentially contributing to the increased potency of inhibin B (a) Inhibins A and B are present in the blood stream as both mature and precursor forms, but differ in that inhibin A appears to be secreted as the precursor aNaC/bA, while inhibin B is secreted as the precursor aC/ ProbB/bB (Fig. 14.1A). Although it appears that these inhibin forms are bioactive, their specific bioactivities have not been established and thus their in vivo activities may differ. (b) The less active 34-kDa diglycosylated forms rather than the 31-kDa monoglycosylated forms of inhibins A and B appear to be the major forms found in the circulation. (c) Other posttranslational modifications (e.g., phosphorylation) may also influence inhibins A and B biological activity. In previous studies (Makanji et al., 2007), we showed that de-glycosylated inhibins A and B exhibit a range of pI forms attributed to other posttranslational changes such as phosphorylation. In other studies, the biological activities of other TGFb ligands, BMP15 and GDF9, are reduced following de-phosphorylation. Phosphorylation of BMP15 and GDF9 was essential for mediation of downstream signaling and de-phosphorylated forms acted as antagonists to their phosphorylated counterparts (McMahon et al., 2008). (d) Our current understanding is that activin B and not activin A is the active agonist at the pituitary in both rats and humans and it is this form which is inhibited by the inhibins. Activin B through its type II
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receptor interacts with both ALK4 and ALK7 type I receptors while activin A operates only through the ALK4 pathway. Is the ALK7 pathway more sensitive to inhibin B interactions?
E. Future directions Regression analysis showed that inhibin B was also an independent regulator of serum LH across the follicular phase of the menstrual cycle. While this relationship has been supported by in vitro studies, there has been little evidence observed in vivo. This was one of the first reports. These studies have also indicated a possible role for AMH in FSH regulation. The regression analysis studies show that AMH is independently and inversely correlated with FSH. There is currently little evidence supporting a pituitary role for AMH; is its role related to its stimulatory action on inhibin synthesis by the ovary potentiating inhibin’s action on the pituitary? Finally, the apparent higher biological activity of inhibin B may not be translated to other biological systems where inhibin A may predominate. For example, in the luteal phase of the menstrual cycle, where inhibin A is in excess of inhibin B, inhibin A may play a greater role in regulating FSH. In conclusion, the regulation of FSH by ovarian inhibins is a multi-step process in which inhibin B appears to be the major inhibin involved. However, it is apparent that structurally different forms of inhibins related to posttranslational changes and other factors such as ovarian steroids and AMH contribute, but these aspects are less well defined.
ACKNOWLEDGMENTS The work was supported by NHMRC (Australia) Project/Program Grant (CAH #494804)/ (DMR #241000) and Research Fellowships (DMR #169201)/(CAH #441125). The authors would also like to thank Prof. Matthew Wilce for the inhibin homology model.
REFERENCES Allendorph, G. P., et al. (2007). BMP-3 and BMP-6 structures illuminate the nature of binding specificity with receptors. Biochemistry 46, 12238–12247. Anderson, R. A. (2001). Clinical studies: Inhibin in the adult male. Mol. Cell. Endocrinol. 180, 109–116. Anderson, R. A., and Sharpe, R. M. (2000). Regulation of inhibin production in the human male and its clinical applications. Int. J. Androl. 23, 136–144. Andersson, A. M., and Skakkebaek, N. E. (2001). Serum inhibin B levels during male childhood and puberty. Mol. Cell. Endocrinol. 180, 103–107. Antenos, M., et al. (2007). N-linked oligosaccharides direct the differential assembly and secretion of inhibin alpha- and betaA-subunit dimers. Mol. Endocrinol. 21, 1670–1684.
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Attardi, B., and Miklos, J. (1990). Rapid stimulatory effect of activin-A on messenger RNA encoding the follicle-stimulating hormone beta-subunit in rat pituitary cell cultures. Mol. Endocrinol. 4, 721–726. Attardi, B., and Winters, S. J. (1993). Decay of follicle-stimulating hormone-beta messenger RNA in the presence of transcriptional inhibitors and/or inhibin, activin, or follistatin. Mol. Endocrinol. 7, 668–680. Attardi, B., et al. (1992). Regulation of FSH beta messenger ribonucleic acid levels in the rat by endogenous inhibin. Endocrinology 130, 557–559. Bedecarrats, G. Y., et al. (2003). Regulation of gonadotropin gene expression by Mullerian inhibiting substance. Proc. Natl. Acad. Sci. USA 100, 9348–9353. Bernard, D. J. (2004). Both SMAD2 and SMAD3 mediate activin-stimulated expression of the follicle-stimulating hormone beta subunit in mouse gonadotrope cells. Mol. Endocrinol. 18, 606–623. Bernard, D. J., et al. (2006). Activin B can signal through both ALK4 and ALK7 in gonadotrope cells. Reprod. Biol. Endocrinol. 4, 52. Bilezikjian, L. M., et al. (1993). Characterization and the regulation of inhibin/activin subunit proteins of cultured rat anterior pituitary cells. Endocrinology 133, 2545–2553. Bilezikjian, L. M., et al. (2004). Autocrine/paracrine regulation of pituitary function by activin, inhibin and follistatin. Mol. Cell. Endocrinol. 225, 29–36. Bilezikjian, L. M., et al. (2006). Pituitary actions of ligands of the TGF-beta family: Activins and inhibins. Reproduction 132, 207–215. Burger, H. (1992). Remembrance: The story of inhibin—The Melbourne version. Endocrinology 131, 1585–1586. Burger, H. G., et al. (1998). Both inhibin A and B respond to exogenous follicle-stimulating hormone in the follicular phase of the human menstrual cycle. J. Clin. Endocrinol. Metab. 83, 4167–4169. Carroll, R. S., et al. (1989). Inhibin, activin, and follistatin: Regulation of follicle-stimulating hormone messenger ribonucleic acid levels. Mol. Endocrinol. 3, 1969–1976. Carroll, R. S., et al. (1991). Activin stabilizes follicle-stimulating hormone-beta messenger ribonucleic acid levels. Endocrinology 129, 1721–1726. Chapman, S. C., and Woodruff, T. K. (2003). Betaglycan localization in the female rat pituitary: Implications for the regulation of follicle-stimulating hormone by inhibin. Endocrinology 144, 5640–5649. Clifton, R. J., et al. (2002). Pachytene spermatocytes in co-culture inhibit rat Sertoli cell synthesis of inhibin beta B-subunit and inhibin B but not the inhibin alpha-subunit. J. Endocrinol. 172, 565–574. Corrigan, A. Z., et al. (1991). Evidence for an autocrine role of activin B within rat anterior pituitary cultures. Endocrinology 128, 1682–1684. Coss, D., et al. (2005). Activin regulates luteinizing hormone beta-subunit gene expression through Smad-binding and homeobox elements. Mol. Endocrinol. 19, 2610–2623. Crofton, P. M., et al. (1997). Changes in dimeric inhibin A and B during normal early puberty in boys and girls. Clin. Endocrinol. (Oxf.) 46, 109–114. Crofton, P. M., et al. (2002a). Dimeric inhibins in girls from birth to adulthood: Relationship with age, pubertal stage, FSH and oestradiol. Clin. Endocrinol. (Oxf.) 56, 223–230. Crofton, P. M., et al. (2002b). Inhibin B in boys from birth to adulthood: Relationship with age, pubertal stage, FSH and testosterone. Clin. Endocrinol. (Oxf.) 56, 215–221. Dalkin, A. C., et al. (1994). Ovarian regulation of pituitary inhibin subunit and activin receptor type II gene expression: Evidence for a nonsteroidal inhibitory substance. Endocrinology 135, 944–949. Dalkin, A. C., et al. (1996). Pituitary activin receptor subtypes and follistatin gene expression in female rats: Differential regulation by activin and follistatin. Endocrinology 137, 548–554.
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de Kretser, D. M., et al. (2002). Inhibins, activins and follistatin in reproduction. Hum. Reprod. Update 8, 529–541. De Winter, J. P., et al. (1996). Truncated activin type II receptors inhibit bioactivity by the formation of heteromeric complexes with activin type I. receptors. Exp. Cell Res. 224, 323–334. di Clemente, N., et al. (1994). Cloning, expression, and alternative splicing of the receptor for anti-Mullerian hormone. Mol. Endocrinol. 8, 1006–1020. Durlinger, A. L., et al. (1999). Control of primordial follicle recruitment by anti-Mullerian hormone in the mouse ovary. Endocrinology 140, 5789–5796. Eldar-Geva, T., et al. (2000). Relationship between serum inhibin A and B and ovarian follicle development after a daily fixed dose administration of recombinant folliclestimulating hormone. J. Clin. Endocrinol. Metab. 85, 607–613. Fraser, H. M., et al. (1999). Follicle-stimulating hormone-inhibin B interactions during the follicular phase of the primate menstrual cycle revealed by gonadotropin-releasing hormone antagonist and antiestrogen treatment. J. Clin. Endocrinol. Metab. 84, 1365–1369. Greenwald, J., et al. (2004). A flexible activin explains the membrane-dependent cooperative assembly of TGF-beta family receptors. Mol. Cell 15, 485–489. Groome, N. P., et al. (1996). Measurement of dimeric inhibin B throughout the human menstrual cycle. J. Clin. Endocrinol. Metab. 81, 1401–1405. Hale, G. E., et al. (2007). Endocrine features of menstrual cycles in middle and late reproductive age and the menopausal transition classified according to the Staging of Reproductive Aging Workshop (STRAW) staging system. J. Clin. Endocrinol. Metab. 92, 3060–3067. Harrington, A. E., et al. (2006). Structural basis for the inhibition of activin signalling by follistatin. EMBO J. 25, 1035–1045. Harrison, C. A., et al. (2004). Modulation of activin and BMP signaling. Mol. Cell. Endocrinol. 225, 19–24. Harrison, C. A., et al. (2005). Antagonists of activin signaling: Mechanisms and potential biological applications. Trends Endocrinol. Metab. 16, 73–78. Hayes, F. J., et al. (2001). Importance of inhibin B in the regulation of FSH secretion in the human male. J. Clin. Endocrinol. Metab. 86, 5541–5546. Hua, V. K., et al. (2008). Effects of protein kinase A and C inhibitors on follicular inhibin and activin during ovulation. Reprod. Biomed. Online 17, 642–651. Itman, C., et al. (2006). All in the family: TGF-beta family action in testis development. Reproduction 132, 233–246. Jaatinen, R., et al. (2002). Activation of the bone morphogenetic protein signaling pathway induces inhibin beta(B)-subunit mRNA and secreted inhibin B levels in cultured human granulosa-luteal cells. J. Clin. Endocrinol. Metab. 87, 1254–1261. Knight, P. G., and Glister, C. (2006). TGF-beta superfamily members and ovarian follicle development. Reproduction 132, 191–206. Lewis, K. A., et al. (2000). Betaglycan binds inhibin and can mediate functional antagonism of activin signalling. Nature 404, 411–414. Ling, N., et al. (1986). A homodimer of the beta-subunits of inhibin A stimulates the secretion of pituitary follicle stimulating hormone. Biochem. Biophys. Res. Commun. 138, 1129–1137. MacConell, L. A., et al. (2002). The distribution of betaglycan protein and mRNA in rat brain, pituitary, and gonads: Implications for a role for betaglycan in inhibin-mediated reproductive functions. Endocrinology 143, 1066–1075. Makanji, Y., et al. (2007). Inhibin A and B in vitro bioactivities are modified by their degree of glycosylation and their affinities to betaglycan. Endocrinology 148, 2309–2316.
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Makanji, Y., et al. (2008). Suppression of inhibin A biological activity by alterations in the binding site for betaglycan. J. Biol. Chem. 283, 16743–16751. Makanji, Y., et al. (2009). Inhibin B is a more potent suppressor of rat follicle-stimulating hormone release than inhibin A in vitro and in vivo. Endocrinology 150, 4784–4793. Mason, A. J., et al. (1996). Characterization and determination of the biological activities of noncleavable high molecular weight forms of inhibin A and activin A. Mol. Endocrinol. 10, 1055–1065. Massague, J., et al. (2005). Smad transcription factors. Genes Dev. 19, 2783–2810. McCullagh, D. R. (1932). Dual endocrine activity of the testes. Science 76, 19–20. McLachlan, R. I., et al. (2002). Effects of testosterone plus medroxyprogesterone acetate on semen quality, reproductive hormones, and germ cell populations in normal young men. J. Clin. Endocrinol. Metab. 87, 546–556. McMahon, H. E., et al. (2008). Phosphorylation of bone morphogenetic protein-15 and growth and differentiation factor-9 plays a critical role in determining agonistic or antagonistic functions. Endocrinology 149, 812–817. Melamed, P. (2010). Hormonal signaling to follicle stimulating hormone beta-subunit gene expression. Mol. Cell. Endocrinol. 314, 204–212. Pernasetti, F., et al. (2001). Cell-specific transcriptional regulation of follicle-stimulating hormone-beta by activin and gonadotropin-releasing hormone in the LbetaT2 pituitary gonadotrope cell model. Endocrinology 142, 2284–2295. Pitteloud, N., et al. (2008). The relative role of gonadal sex steroids and gonadotropinreleasing hormone pulse frequency in the regulation of follicle-stimulating hormone secretion in men. J. Clin. Endocrinol. Metab. 93, 2686–2692. Plant, T. M., and Marshall, G. R. (2001). The functional significance of FSH in spermatogenesis and the control of its secretion in male primates. Endocr. Rev. 22, 764–786. Robertson, D. M., et al. (2008). A proposed classification system for menstrual cycles in the menopause transition based on changes in serum hormone profiles. Menopause 15, 1139–1144. Robertson, D. M., et al. (2009). Interrelationships between ovarian and pituitary hormones in ovulatory menstrual cycles across reproductive age. J. Clin. Endocrinol. Metab. 94, 138–144. Sealfon, S. C., et al. (1990). Hormonal regulation of gonadotropin-releasing hormone receptors and messenger RNA activity in ovine pituitary culture. Mol. Endocrinol. 4, 1980–1987. Setchell, B. P., and Jacks, F. (1974). Inhibin-like activity in rete testis fluid. J. Endocrinol. 62, 675–676. Shi, Y., and Massague, J. (2003). Mechanisms of TGF-beta signaling from cell membrane to the nucleus. Cell 113, 685–700. Shi, F. T., et al. (2009). Effects of endogenous growth differentiation factor 9 on activin A-induced inhibin B production in human granulosa-lutein cells. J. Clin. Endocrinol. Metab. 94, 5108–5116. Tebar, M., et al. (2000). Regulation of inhibin/activin subunits and follistatin mRNA expression in the rat pituitary at early estrus. Life Sci. 67, 2549–2562. Thackray, V. G., et al. (2010). Hormones in synergy: Regulation of the pituitary gonadotropin genes. Mol. Cell. Endocrinol. 314, 192–203. Thompson, T. B., et al. (2003). Structures of an ActRIIB:activin A complex reveal a novel binding mode for TGF-beta ligand:receptor interactions. EMBO J. 22, 1555–1566. Thompson, T. B., et al. (2004). Beta A versus beta B: Is it merely a matter of expression? Mol. Cell. Endocrinol. 225, 9–17. Tsuchida, K., et al. (2004). Activin isoforms signal through type I receptor serine/threonine kinase ALK7. Mol. Cell. Endocrinol. 220, 59–65.
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Uccella, S., et al. (2000). Localization of inhibin/activin subunits in normal pituitary and in pituitary adenomas. Pituitary 3, 131–139. Visser, J. A., et al. (2006). Anti-Mullerian hormone: A new marker for ovarian function. Reproduction 131, 1–9. Weiss, J., et al. (1995). Transcriptional activation of the follicle-stimulating hormone betasubunit gene by activin. Endocrinology 136, 1885–1891. Welt, C. K., and Schneyer, A. L. (2001). Differential regulation of inhibin B and inhibin a by follicle-stimulating hormone and local growth factors in human granulosa cells from small antral follicles. J. Clin. Endocrinol. Metab. 86, 330–336. Welt, C. K., et al. (2001). Differential regulation of inhibin A and inhibin B by luteinizing hormone, follicle-stimulating hormone, and stage of follicle development. J. Clin. Endocrinol. Metab. 86, 2531–2537. Welt, C. K., et al. (2003). Control of follicle-stimulating hormone by estradiol and the inhibins: Critical role of estradiol at the hypothalamus during the luteal-follicular transition. J. Clin. Endocrinol. Metab. 88, 1766–1771. Wiater, E., et al. (2009). Endogenous betaglycan is essential for high-potency inhibin antagonism in gonadotropes. Mol. Endocrinol. 23, 1033–1042. Woodruff, T. K., and Mather, J. P. (1995). Inhibin, activin and the female reproductive axis. Annu. Rev. Physiol. 57, 219–244. Xia, Y., and Schneyer, A. L. (2009). The biology of activin: Recent advances in structure, regulation and function. J. Endocrinol. 202, 1–12. Yamada, Y., et al. (2004). Differential activation of the luteinizing hormone beta-subunit promoter by activin and gonadotropin-releasing hormone: A role for the mitogenactivated protein kinase signaling pathway in LbetaT2 gonadotrophs. Biol. Reprod. 70, 236–243.
C H A P T E R
F I F T E E N
Activin A in Nonalcoholic Fatty Liver Disease Arne Yndestad,* John Willy Haukeland,† Tuva B. Dahl,* Bente Halvorsen,*,‡ and Pa˚l Aukrust*,‡,§ Contents I. Introduction II. Nonalcoholic Fatty Liver Disease A. Natural history, diagnosis, and epidemiology B. Pathophysiology of NAFLD C. NAFLD and metabolic syndrome D. Mechanisms for hepatic lipid accumulation in NAFLD E. Apoptosis and fibrosis—Important processes in progressive NAFLD F. NAFLD and systemic inflammation III. Activin A in Liver Biology and Pathology A. General properties of activin A B. Regulation of activin A bioactivity IV. Activin A in NAFLD A. Activin A in hepatic disorders B. Activin A in patients with NAFLD C. Activin A and the metabolic syndrome D. Activin A and apoptosis E. Activin A and hepatic fibrosis F. Activin A and inflammation in NAFLD V. Conclusion and Future Perspectives References
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Abstract Nonalcoholic fatty liver disease (NAFLD) is emerging as one of the most common causes of abnormal liver function. NAFLD is a spectrum of disease ranging from simple steatosis (i.e., fat accumulation) of the liver to nonalcoholic * Research Institute for Internal Medicine, Oslo University Hospital Rikshospitalet, Oslo, Norway Department of Gastroenterology, Oslo University Hospital Aker, Oslo, Norway Faculty of Medicine, University of Oslo, Oslo, Norway } Section of Clinical Immunology and Infectious Diseases, Oslo University Hospital Rikshospitalet, Oslo, Norway { {
Vitamins and Hormones, Volume 85 ISSN 0083-6729, DOI: 10.1016/B978-0-12-385961-7.00015-9
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2011 Elsevier Inc. All rights reserved.
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steatohepatitis (NASH) with inflammation and fibrosis. NAFLD involves the accumulation of triglycerides in hepatocytes, necrosis, and apoptosis of these cells, accompanied by inflammatory and fibrogenic responses within the liver, potentially leading to the development of cirrhosis. Activin A, a member of the transforming growth factor-b family of cytokines, has been recognized as a multifunctional cytokine expressed in a wide range of cells and tissues with roles in regulation of wound repair, cell differentiation, apoptosis, and inflammation. Growing evidence suggests that activin A could be involved in the pathogenesis of various liver disorders such as acute liver injury, chronic viral hepatitis, and certain hepatic malignancies, and recently we demonstrated an involvement of activin A in NAFLD. In this chapter, after a general introduction to NAFLD and activin A biology, we elaborate a potential pathogenic role of activin A in the development and progression of NAFLD. ß 2011 Elsevier Inc.
I. Introduction Nonalcoholic fatty liver disease (NAFLD) is emerging as one of the most common causes of abnormal liver function, and in the Western world the estimated prevalence is reported to be about 20% (Bjornsson and Angulo, 2007). Histologically, NAFLD is a spectrum of disease ranging from simple steatosis (i.e., fat accumulation) of the liver to nonalcoholic steatohepatitis (NASH) with inflammation and fibrosis and subsequently, extensive fibrosis and NASH-associated cirrhosis characterizing the most advanced forms of NASH. While NASH implies a risk of progressive liver disease (Ekstedt et al., 2006), simple steatosis might be regarded as a benign condition (Dam-Larsen et al., 2004). NAFLD involves the accumulation of triglycerides in hepatocytes, necrosis, and apoptosis of these cells, accompanied by inflammatory and fibrogenic responses within the liver, potentially leading to the development of cirrhosis. The two-hit model summarizes the important early metabolic events leading to fat accumulation and subsequently hepatocellular necrosis and inflammation in NASH (Day and James, 1998). Based on the two-hit model, it is of major importance to identify factors that could trigger hepatic fat accumulation as well as mediators that could promote the hepatic transition from simple steatosis to NASH. The metabolic syndrome with obesity, dyslipidemia, and insulin resistance (IR) is frequently associated with NAFLD (Marchesini et al., 2003). Although these conditions, as well as inflammation and oxidative stress (Seki et al., 2002), may predispose to NAFLD development, the mechanisms that underlie hepatic fat accumulation and triggering of hepatocyte injury and hepatic fibrosis in NASH are still largely unknown. In particular, little is known about the mediators
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that could trigger the extensive hepatic fibrogenic response in certain individuals with NAFLD, leading to advanced NASH. Activin A is a member of the transforming growth factor (TGF)-b superfamily (de Caestecker, 2004) and was originally described as an inducer of follicle-stimulating hormone release (Ling et al., 1986). More recently, activin A has been recognized as a multifunctional cytokine expressed in a wide range of cells and tissues with roles in regulation of wound repair, cell differentiation, apoptosis, and inflammation, and growing evidence implicates activin A in the pathogenesis of various inflammatory disorders such as rheumatoid arthritis, inflammatory bowel disease, and atherosclerosis (Werner and Alzheimer, 2006). Studies also suggest that activin A could be involved in the pathogenesis of various liver disorders such as acute liver injury, chronic viral hepatitis, and certain hepatic malignancies (RodgarkiaDara et al., 2006), and recently we demonstrated an involvement of activin A in NAFLD (Yndestad et al., 2009). In this chapter, after a general introduction to NAFLD and activin A biology, we elaborate a potential pathogenic role of activin A in the development and progression of NAFLD.
II. Nonalcoholic Fatty Liver Disease A. Natural history, diagnosis, and epidemiology For decades, it has been known that obesity is associated with fatty liver (Zelman, 1952). However, the scientific interest for this observation was scarce until 1980 when Ludwig introduced the term NASH in 20 patients with no or only occasional intake of alcohol (Ludwig et al., 1980). During recent years, it has been common to use the term NAFLD for a wide spectrum of liver damage, ranging from simple steatosis of the liver to NASH and cirrhosis on the basis of hepatic and/or systemic responses to long-standing fatty infiltration of the liver (Angulo, 2002). Even hepatocellular carcinoma is now recognized as a possible complication of NAFLD (Bugianesi et al., 2002). The natural history of NAFLD is strongly dependent on its stage. It is generally believed that patients with simple steatosis carry a minimal risk of progression to cirrhosis (Dam-Larsen et al., 2004; Teli et al., 1995), while having NASH implies a risk of progressive fibrosis with subsequent development of cirrhosis and hepatic failure (Adams et al., 2005; Fassio et al., 2004; Harrison et al., 2003). In a large follow-up study, Ekstedt et al. found that 10% of patients with NASH developed end-stage liver disease after median 13.7 years (Ekstedt et al., 2006). Furthermore, compared with a matched reference population, both overall mortality and mortality due to liver disease were significantly increased among patients with NASH, but not in patients with simple steatosis.
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Cross-sectional studies based on ultrasonography have shown NAFLD to be a common disease, affecting 10–20% of the general adult population in both Western and developing countries (Lewis and Mohanty, 2010). By use of magnetic resonance spectroscopy, an even higher estimate (31%) was found in the general adult population in the US (Browning et al., 2004), probably reflecting the high prevalence of obesity and metabolic syndrome in this country. Less is known about the prevalence of NASH. A frequency of 2.7% in lean patients found at autopsy has often been cited (Wanless and Lentz, 1990). Recently, NASH was reported in 2.2% of potential living donors in Korea (Lee et al., 2007). Although data from transplantation registries clearly show that NASH indeed is a cause of cirrhosis, the incidence rate of cirrhosis due to NASH in the general population is not known.
B. Pathophysiology of NAFLD Molecular mechanisms underlying hepatic steatosis, and in some cases, the development of liver injury, inflammation, fibrosis, and NASH have been an area of intensive research during the recent years (Browning and Horton, 2004). A fundamental feature in the disease process is the accumulation of fat within the hepatocytes. This event represents the “first hit” as conceptualized by Day in 1998 (Day and James, 1998). The condition may progress to NASH as a result of “second hits,” such as oxidative stress, lipid peroxidation, endoplasmatic stress, and activation of inflammatory and profibrotic cytokines (Cortez-Pinto et al., 2006). Accumulation of fibrous tissue will then depend on the balance between fibrogenesis and remodeling of extracellular matrix (ECM) as well as the degree of hepatocyte apoptosis or necrosis (Iredale, 2007). These processes may ultimately result in cirrhosis, in which case the degree of fat accumulation often is reduced, and possibly may have disappeared. The causes of the pathological processes in NAFLD are complex. In the gross majority, overweight and a sedentary lifestyle are thought to be essential in the development of NAFLD (Angulo, 2002). Partly depending on genetic susceptibility, it induces IR which may promote fat accumulation in the liver being part of the “first hits.” The mediators that trigger the development from simple steatosis (“second hits”) are less clear but seem to involve inflammatory cytokines, oxidative stress, and fibrogenic mediators. These mechanisms will be discussed in more details below.
C. NAFLD and metabolic syndrome NAFLD is seen much more frequently in patients with overweight and/or features of the metabolic syndrome. In all, 76% of obese patients (Bellentani et al., 2000), 49% of patients with type 2 diabetes mellitus (T2DM)
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(Gupte et al., 2004), and 31% of patients with arterial hypertension (Donati et al., 2004) have been found to have NAFLD. Moreover, according to pooled data of histological results in 12 different studies on patients undergoing bariatric surgery, the prevalence of NAFLD and NASH were 91% and 37%, respectively (Machado et al., 2006). IR, hyperglycemia, hyperinsulinemia, and altered lipid metabolism may be a common link between these clinical conditions and NAFLD. Albeit various definitions of the metabolic syndrome, they are all based upon criteria related to (i) elevated blood pressure, (ii) overweight or abdominal adiposity, (iii) elevated plasma triglycerides and low levels of high-density lipoprotein (HDL)-cholesterol, and (iv) disturbed glucose homeostasis defined as either impaired fasting glucose, impaired glucose tolerance, or IR (Day, 2007). IR has been clearly demonstrated in nondiabetic patients with NAFLD by use of the euglycemic hyperinsulinemic clamp technique and the frequently sampled intravenous glucose tolerance test (Bugianesi et al., 2005; Pagano et al., 2002; Sanyal et al., 2001). Furthermore, numerous crosssectional studies have revealed high prevalence of the metabolic syndrome and/or T2DM in patients with NAFLD (Marchesini et al., 2003; Pagano et al., 2002). Thus, the presence of IR in patients with NAFLD and the increased prevalence of related clinical conditions have lead many authors to suggest NAFLD be perceived part of the metabolic syndrome (Loria et al., 2005).
D. Mechanisms for hepatic lipid accumulation in NAFLD Insulin regulates lipid and glucose metabolism primarily through its action on adipose tissue, muscles, and liver. The overall effect of insulin on energy homeostasis is to promote energy uptake in peripheral tissues (i.e., muscles and adipose tissue). The circulating amounts of glucose and free fatty acids (FFA) are thereby kept within certain limits, both postprandially and in fasting conditions. However, in IR states, blood levels of these substrates may increase as well as levels of glycerol and triglycerides. These changes together with the compensatory hyperinsulinemia typically found in early stages of IR, may promote de novo lipogenesis and fat accumulation in the liver (Donnelly et al., 2005). The concept of hepatic IR is not always properly defined. While there is resistance to the effect of insulin regarding glucose homeostasis (i.e., gluconeogenesis is not inhibited as normal), the normal lipogenic response in the liver upon stimulation with insulin (Horton et al., 2002) may be intact (Lind, 2004; Shimomura et al., 2000). Interestingly, insulin’s ability to suppress hepatic glucose production is reduced in fatty liver, thus illustrating an important interrelationship between lipid and glucose metabolism with insulin as a common regulator (Seppala-Lindroos et al., 2002).
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E. Apoptosis and fibrosis—Important processes in progressive NAFLD While accumulation of fat is regarded as the “first hit” in NAFLD pathogenesis, disease progression toward NASH and cirrhosis is postulated to result from “second hits” that lead to hepatocyte injury and death, activation of hepatic stellate cells (HSCs) and fibrosis, and inflammation. Potential “second hits” may be oxidative stress, lipid peroxidation, endoplasmatic stress, and activation of inflammatory and profibrotic cytokines (CortezPinto et al., 2006). Intrahepatic FFAs are hepatotoxic in several ways and are potentially responsible for the generation of many of these “second hits” ( Jou et al., 2008). First, they can be directly cytotoxic via lipoapoptosis (Unger and Orci, 2002). Second, FFAs may activate nuclear factor-kB and induce several inflammatory cytokines such as tumor necrosis factor (TNF)a (see below) (Tilg and Moschen, 2008). Finally, FFA metabolism leads to the generation of reactive oxygen species (Tilg and Moschen, 2008). Oxidative stress may have detrimental effects on cellular function, including mitochondrial damage, and may also cause the formation of highly toxic lipid peroxides (Tilg and Moschen, 2008). A common ultimate consequence of these processes may be hepatocyte death, both through necrosis and in particular through apoptosis. The above mechanisms will also lead to activation of HSCs and increased generation of ECM components. This is further stimulated by loss of hepatocytes that will lead to deposition of ECM as a normal healing response. Apoptosis and fibrosis are also potentially tightly linked processes. Furthermore, apoptotic hepatocytes may be ingested by Kupffer cells and HSC which subsequently release fibrogenic cytokines such as TGF-b (Canbay et al., 2003; Fadok et al., 1998). Importantly, TGF-b may again also induce hepatocyte apoptosis (Gressner et al., 1997), suggesting that the interaction between hepatic apoptosis and fibrosis may represent a pathogenic loop in the progression of NAFLD.
F. NAFLD and systemic inflammation Over the last decade, numerous studies have shown an association between conditions related to IR and inflammatory mediators (Dandona et al., 2005). Even in healthy individuals, a consistent relationship between C-reactive protein (CRP) and markers of IR has been demonstrated (Festa et al., 2000). Furthermore, an increasing amount of basic research has revealed considerable cross-talk between inflammatory pathways and insulin signaling. Accordingly, systemic inflammation seems to play an important pathogenic role in conditions related to IR (Wellen and Hotamisligil, 2005). This includes NAFLD and several studies have shown elevated circulating levels of inflammatory cytokines such as
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TNFa, interleukin (IL)-6 and monocyte chemoattractant protein (MCP)-1 in NAFLD patients with particularly enhanced levels in patients with NASH and cirrhosis (Haukeland et al., 2006; Tilg and Moschen, 2008). Importantly, inflammation and inflammatory cytokines may affect NAFLD pathogenesis at several levels. TNFa, originally named cachectin for its ability to cause weight loss in animal models (Cerami and Beutler, 1988), has, for example, potent effects on insulin sensitivity and lipid and glucose metabolism (Beier et al., 1992; Uysal et al., 1997), and treatment with neutralizing anti-TNFa antibodies in rats with IR have been shown to improve hepatic steatosis and fibrosis (Tilg and Moschen, 2008). Thus, inflammation and TNFa cannot only be related to cachexia but also to obesity. Hence, several studies have shown that adipose tissue may contribute to low-grade systemic inflammation and vice versa, inflammation may promote dysfunction of adipocytes and adipose tissue and by its ability to inhibit lipoprotein lipase, TNFa may induce hypertriglyceridemia (Hube and Hauner, 1999). As for MCP-1, this chemokine has been found to induce macrophage recruitment into adipose tissue in obese subjects (Kanda et al., 2006). Recently, Weisberg et al. gave further support to this finding reporting reduced inflammatory profile and macrophage content in adipose tissue, improved insulin sensitivity, and even marked reduction of hepatic steatosis in obese mice deficient of CCR2 (receptor for MCP-1) (Weisberg et al., 2006), further underscoring a link between inflammation, metabolic syndrome, and hepatic steatosis.
III. Activin A in Liver Biology and Pathology A. General properties of activin A Activin A is a member of the TGF-b superfamily, a large family of over 30 structurally related proteins (Piek et al., 1999). TGF-b superfamily cytokines play pivotal roles in regulation of tissue homeostasis, organ development, inflammation, cell proliferation, and apoptosis (de Caestecker, 2004; Ling et al., 1986). In fact, activin A has been recognized as a multifunctional cytokine expressed in a wide range of tissues and cells, and growing evidence implicates activin A in the pathogenesis of a variety of disorders ranging from rheumatoid arthritis, bone disorders, sepsis, inflammatory bowel disease, atherosclerosis, and chronic heart failure to certain malignancies (Chen et al., 2002; Smith et al., 2004; Werner and Alzheimer, 2006; Yndestad et al., 2004). Activin A signal transduction is similar to that of other TGF-b superfamily members (Pangas and Woodruff, 2000). Binding of activin A to the activin type II receptors (ActR) leads to the recruitment of a type I receptor, ActRIB/activin receptor-like kinase (ALK)4, and formation of a heteromeric
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receptor complex (Fig. 15.1). This activated activin receptor complex phosphorylates the receptor-associated Smads, Smad2 or Smad3, that subsequently can form a heterocomplex with the co-Smad, Smad4. The resulting Smad complex will translocate into the nucleus and, in collaboration with transcriptional coactivators or corepressors (e.g., CBP/p300), regulate the expression of the target genes. In addition, activin A may also activate Smad-independent signaling pathway, such as the mitogen-activated protein kinases p38 and JNK (Rodgarkia-Dara et al., 2006).
B. Regulation of activin A bioactivity The activity and response to activin A are tightly modulated at several levels and this has been reviewed elsewhere (Chen et al., 2002; Rodgarkia-Dara et al., 2006). The influence of extracellular ligand-binding proteins on Activin A
Follistatin Follistatin-like 3
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Figure 15.1 Activin A signal transduction and regulation of activity. The figure illustrates the activation of the Smad pathway by activin A. Binding of activin A leads to the formation and phosphorylation of a heteromeric receptor complex that recruits and phosphorylates receptor-associated Smads, that is, Smad2 or Smad3. These form a complex with Smad4 that translocate into the nucleus and, in collaboration with transcriptional coactivators or corepressors (e.g., CBP/p300), regulate the expression of the target genes. The bioactivity of activin A is regulated at several levels. This includes the binding and inactivation of activin A by follistatin or follistatin-like 3. Intracellularly, the activity of activin A may be limited by Smad7 that inhibits the formation of an activated Smad complex.
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activin A activity may be of particular importance. Follistatin and follistatinlike 3 (FSTL3) are the best characterized of these binding proteins and prevent activin A, and also other TGF-b superfamily members from interacting with its signaling receptors (Fig. 15.1). It has been shown that follistatin binds activin A with nearly irreversible kinetics, indicating that activin A is practically inactivated upon binding. Follistatin and FSTL3 are expressed in many of the same tissues as activin A and, importantly, activin A may induce the synthesis of these binding proteins suggesting that they represent a negative feedback loop to restrict activin A bioactivity (Chen et al., 2002; Rodgarkia-Dara et al., 2006). The Smad2/3 pathway represents the principal effector of activin A signaling. Importantly, this pathway may be inhibited by the negative regulator Smad7 (Fig. 15.1). Smad7 binds tightly to activated type I receptors (see above) and prevents binding of other Smads to the receptors (Chen et al., 2002). Smad7 expression is induced by activin A, and Smad7mediated inhibition of activin A signaling therefore represents another negative feedback mechanism. Importantly, Smad7 also represents a site where signals from other pathways converge, and interferon-g, TNFa, and IL-1 have all been shown to increase Smad7 expression, potentially leading to inhibited activin A signaling (Rodgarkia-Dara et al., 2006).
IV. Activin A in NAFLD A. Activin A in hepatic disorders Experimental studies have shown that activin A has a range of effects on hepatic cells that may support a pathogenic role for this cytokine in hepatic disorders (Rodgarkia-Dara et al., 2006). These include induction of hepatocyte apoptosis and inhibition of hepatocyte growth (Schwall et al., 1993). However, activin A has been shown to promote ECM production in HSCs and tubulogenesis of sinusoidal endothelial cells, and thus contributes to restoration of tissue architecture during liver regeneration (De Bleser et al., 1997; Patella et al., 2006). Clinical studies suggest an involvement of activin A in the pathogenesis of a range of liver disorders (Rodgarkia-Dara et al., 2006). Serum levels of activin A are increased in patients with chronic viral hepatitis and alcohol-induced cirrhosis (Patella et al., 2001; Yuen et al., 2002), as well as in patients acute liver failure (Hughes and Evans, 2003). Notably, it seems like the ratio between follistatin and activin A in blood may provide important prognostic information in the latter disorder (Lin et al., 2006). Furthermore, increased serum levels of activin A have been found in patients with hepatocellular carcinoma and it has been suggested that deregulation of the growth inhibitory control and apoptotic effect of activin A on hepatocytes might promote tumor development (Deli et al., 2008).
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Finally, activin A and follistatin has also attracted interest with regards to a regulatory role in liver regeneration in response to partial hepatectomy or liver injury (Rodgarkia-Dara et al., 2006).
B. Activin A in patients with NAFLD Results in our studies indicate that activin A signaling may be of pathogenic importance also in patients with NAFLD (Yndestad et al., 2009). First, serum levels of activin A and follistatin are increased in patients with NAFLD, and particularly in patients with NASH. Second, we found that the hepatic gene expression of follistatin was significantly decreased in patients with NAFLD so that the activin A/follistatin ratio actually is increased. Since the activity of activin A in various tissues is modulated by its endogenous inhibitor follistatin which binds activin A with high affinity and neutralizes its biological action (see above) (Rodgarkia-Dara et al., 2006), the increased activin A/follistatin ratio observed in our study may suggest an important role for activin A-mediated effects at the cellular level in the liver. Immunohistochemical staining revealed that both activin A and follistatin were expressed in hepatocytes, biliary epithelial cells, and occasionally endothelial cells in portal areas. Although hepatocytes constitute about 80% of total cell mass within the liver, identification of the exact cell type responsible for the altered hepatic gene expression observed in our study is difficult to know. Moreover, the explanation for elevated serum levels of activin A is not clear from our data, as the hepatic gene expression actually tended to be reduced. Extrahepatic secretion of activin A may provide one possible explanation. Notably, systemic activin A levels is elevated in disorders that may share pathogenic mechanisms, such as coronary artery disease (Smith et al., 2004). Whatever the mechanisms for the enhanced activin A levels in NAFLD, with increased serum levels and enhanced activin A/follistatin ratio in the liver potentially reflecting increased activin A bioactivity, this disturbed activin A levels could potentially contribute to the pathogenesis of NAFLD. Thus, although the pathogenesis of NAFLD-related disease is incompletely understood, metabolic disturbances, hepatocyte injury and death, fibrosis, and inflammation seem to be important features, and notably, activin A could potentially modulate all these interacting pathogenic processes (Fig. 15.2).
C. Activin A and the metabolic syndrome As mentioned above, NAFLD may be regarded as the hepatic manifestation of the metabolic syndrome involving obesity, IR and diabetes, dyslipidemia, and hypertension (Day, 2007; Jou et al., 2008). In the multihit hypotheses of NAFLD pathogenesis, IR is thought to initiate the first “hit,” steatosis. The steatosis develops partly as a result of increased levels of FFA released from
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Normal liver Activin A Insulin sensitivity Glucose metabolism Fatty acid metabolism
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Figure 15.2 Activin A and the two-hit hypothesis for pathogenesis of nonalcoholic fatty liver disease (NAFLD). The first hit, insulin resistance and metabolic disturbances, results in hepatic lipid accumulation, producing steatosis. The second hits, for example, oxidative stress and inflammatory mediators, lead to progressive NAFLD with loss of hepatocytes, hepatic stellate cell activation, and fibrosis and inflammation. This results in nonalcoholic steatohepatitis (NASH). Activin A may affect NAFLD pathogenesis in several ways. Its effects on metabolism are potentially beneficial and may counteract the first hit and lipid accumulation. In contrast, activin A has effects on apoptosis and fibrosis that may contribute to progressive NAFLD.
muscle and adipose tissue. These are taken up in the liver, are incorporated into triglycerides, or are oxidized. Several lines of evidence suggest that activin A may contribute to the disturbed glucose and lipid metabolism in NAFLD. 1. Activin A and hepatic FA metabolism We showed that activin A has potentially important effects on FA metabolism in hepatocytes (Yndestad et al., 2009). Intrahepatic accumulation of triglycerides, at least partly involving enhanced de novo FA synthesis in the liver (Donnelly et al., 2005), is a key event in the development of steatohepatitis. This accumulation of lipids is counteracted by increased mitochondrial b-oxidation leading to a compensatory effect with increased removal of FA (Fromenty et al., 2004). We reported that activin A potently suppresses fatty acid synthase (FAS) and enhances carnitine palmitoyltransferase (CPT)-I activity in hepatocytes, enzymes that are of major importance for hepatic lipogenesis and mitochondrial b-oxidation of FA,
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respectively. In fact, CPT-I is considered rate controlling of mitochondrial b-oxidation through different mechanisms (Kerner and Hoppel, 2000). Activin A could therefore have favorable effects on lipid metabolism in the liver in relation to NAFLD by enhancing FA oxidation and suppressing de novo FA synthesis. The relevance of these in vitro findings, however, may be limited as the cells in the study were not a model of fat-filled hepatocytes. We also found that these activin A-mediated effects on the synthesis and oxidation of FA were accompanied by several changes in FA composition such as decreased levels of saturated FA, increased levels of monosaturated FA, and altered composition within the n-3 and n-6 polyunsaturated FA (PUFA) (e.g., increased weight% of eicosapentaenoic acid (EPA)). While some of these changes have been reported in experimental steatohepatitis (i.e., increased ratio of monounsaturated:saturated FA) (Larter et al., 2008), the increase in EPA could represent a potential favorable effect of activin A in NAFLD. In fact, some recent studies have suggested beneficial effects of EPA supplementation in NASH patients (Tanaka et al., 2008). If some of the activin A-mediated effects on lipid metabolism in hepatocytes also are operating in vivo within the liver of NAFLD patients, this cytokine could counteract lipid accumulation in these patients. 2. Activin A and glucose metabolism In addition to the modulatory effects on FA and lipid metabolism, activin A may also influence glucose metabolism. Thus, activin A plays an important role in pancreas development and islet b-cell maturation (Yamaoka et al., 1998) and has also been shown to be important in the regeneration of b-cells and improvement of glucose metabolism in a rat model of streptozocininduced diabetes (Li et al., 2004). These stimulatory effects of activin A on pancreatic b-cell differentiation seem to, at least in part, be mediated by activation of PAX4, one of the major transcription factors that are involved in this process (Ueda, 2000). Importantly, activin A may also directly induce insulin secretion independently of glucose levels at least partly by stimulating Ca2þ entry into islet cells (Furukawa et al., 1995). These studies may all suggest a favorable effect of activin A in NAFLD. A recent study by Mukherjee et al. (2007) may lead to the opposite conclusion. These authors showed that mice deficient in follistatin-like 3, a potent activin A and also myostatin antagonist, developed increased pancreatic islet number and size, b-cell hyperplasia, decreased visceral fat mass, improved glucose tolerance, and enhanced insulin sensitivity; they also, somewhat surprisingly, developed hepatic steatosis (Mukherjee et al., 2007). The reason for this latter finding is not clear. Nonetheless, these findings underscore the complex effect of activin A on glucose and lipid metabolism as well as the multifaceted interplay between these two metabolic pathways.
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D. Activin A and apoptosis Apoptosis potentially represents an important mechanism for the loss of hepatocytes that occurs during the progression of NAFLD to NASH and further to cirrhosis. Regulation of life and death is a central feature in activin A biology, and activin A has shown to both induce and inhibit apoptosis depending on cell type and cell milieu (Chen et al., 2002). Already 17 years ago, it was shown that subcutaneously administered activin A in rats induces apoptosis in hepatocytes (Schwall et al., 1993), and interestingly, apoptotic changes were predominantly observed in the centrilobular region of the liver, the same area that is typically involved in NAFLD (Yeh and Brunt, 2007). Furthermore, administration of the activin-binding protein follistatin in an animal model of liver fibrosis has been shown to reduce fibrosis development, at least partly due to inhibited hepatocyte apoptosis (Patella et al., 2006). Several studies have shown that activin A may induce apoptosis in various cancer cells involving activation of caspases and downregulatory effect of anti-apoptotic mediators such as bcl-2. Dysregulation of activin A activity has also been suggested to play a role in hepatic carcinogenesis (Grusch et al., 2006). Neoplastic hepatocytes are very sensitive to activin A-induced apoptosis (Grusch et al., 2006), and increased inactivation by follistatin and therefore reduced apoptosis might very well represent a key carcinogenic mechanism. Thus, it can be speculated that increased activin A signaling in NAFLD could act as a pro-apoptotic stimulus in hepatocytes already vulnerable due to other causes such as lipotoxicity and oxidative stress.
E. Activin A and hepatic fibrosis Liver fibrogenesis can be considered a wound-healing response to chronic liver injury and hepatocyte loss. The principal cell type responsible for production of excessive ECM in the liver is the HSC. Upon activation, the HSC changes phenotype from quiescent vitamin A storing cells into myofibroblast-like cells, acquiring contractile, inflammatory, and fibrogenic properties (Bataller and Brenner, 2005). The activation of the HSC occurs as a result of several stimuli, including cytokines released from damaged hepatocytes and activin A could clearly also be involved in this process. The control of tissue homeostasis involving regulation of the quantity and quality of ECM is a fundamental property of several members of the TGF-b superfamily, particularly TGF-b itself and activin A (Hubner et al., 1999). These regulatory effects on ECM and fibrogenesis seem also to be involved in the proposed role of activin A in the pathogenesis of various liver disorders (Rodgarkia-Dara et al., 2006). In addition to in NAFLD, elevated levels of activin A have been found in liver disorders where fibrosis is important, such as alcohol-induced cirrhosis and chronic viral hepatitis
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(Patella et al., 2001; Yuen et al., 2002), as well as in the liver in experimental models of liver fibrosis (De Bleser et al., 1997; Gold et al., 2003). Importantly, administration of the activin A–antagonist follistatin to CCl4-treated rats markedly reduced the development of hepatic fibrosis most probably via inhibited activin A-induced HSC activation (Patella et al., 2006). In relation to activin A and NAFLD-related fibrosis, the suggested complex interplay between the hepatocytes and the HSC involving paracrine and autocrine signaling may be relevant (Werner and Alzheimer, 2006). Both cell types may secrete activin A, but while the response in the HSC is activation, the hepatocytes are pushed toward apoptosis. The finding of significantly increased serum levels of activin A in NASH patients with advanced fibrosis as compared to those with no or mild fibrosis fits well with the hypothesis that activin A plays a role in liver fibrogenesis (Yndestad et al., 2009). While previously considered an irreversible one-directional event, current understanding of liver fibrogenesis describes it as a dynamic process not only involving production of ECM but also modification and resolution of ECM by matrix metalloproteinases (MMP) which are kept under control by tissue inhibitor of MMPs (TIMPs) (Bataller and Brenner, 2005; Han, 2006; Iredale, 2007). When stimulated with activin A, the secretion of MMPs increased significantly in hepatocytes in our study (Yndestad et al., 2009). This effect could potentially represent an anti-fibrotic effect. However, although MMPs from hepatocytes could promote resolution of fibrosis, the net result of activin A signaling in the liver could still be increased fibrogenesis, since the HSC becomes activated when exposed for activin A. Besides, recent in vitro studies on primary rat hepatocytes have shown that activin A may induce expression of connective tissue growth factor which would promote fibrogenesis (Gressner et al., 2008). Moreover, we showed that activin A increased the release of TGF-b from hepatocytes, further strengthening a role for activin A in the promotion of fibrogenesis in NAFLD-related disease.
F. Activin A and inflammation in NAFLD Inflammatory cytokines may play a pathogenic role in the development and progression of NAFLD as outlined above (Tilg and Moschen, 2008). Activin A is a cytokine with potent regulatory properties on inflammation and both pro- and anti-inflammatory effects have been described ( Jones et al., 2004). In macrophages, activin A has been shown to potently stimulate the production of TNFa and IL-1b, as well as the expression of cyclooxygenase-2 (COX-2) and inducible nitric oxide synthase (iNOS) ( Jones et al., 2004). However, the effect of activin A on macrophages within the liver (i.e., Kupffer cells) has yet to be examined. Moreover, activin A can have important effects on the recruitment of leukocytes into
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inflamed tissue and has been shown to have chemotactic effects on monocytes ( Jones et al., 2004). The anti-inflammatory cytokine, IL-10, has been suggested to be protective in NAFLD (den Boer et al., 2006), and activin A has previously been shown to oppose the actions of IL-10 in a prostatic epithelial cell line (Wang et al., 1999). Thus, activin A clearly possesses properties that can indicate a role in increasing inflammation in NAFLD disease progression. However, anti-inflammatory actions of activin A are also well documented and activin A has been reported to inhibit IL-1 and IL-6 activity by both inhibiting their production and antagonizing their action ( Jones et al., 2004). In relation to NAFLD, the ability of activin A to antagonize IL-6-mediated effects in hepatocytes is potentially of pathogenic relevance (Russell et al., 1999). The role of activin A in regulating the inflammatory responses in NAFLD or other hepatic disorders has not been addressed directly. Based on the dichotomy between pro- and anti-inflammatory actions of activin A, it is practically impossible to predict in which direction activin A pushes the inflammation in NAFLD. However, we have previously shown that activin A has anti-inflammatory effects on peripheral blood mononuclear cells (PBMC) from patients with coronary heart disease characterized by persistent low-grade systemic inflammation (Smith et al., 2004). In contrast, this cytokine markedly enhanced the release of inflammatory cytokines in PBMCs from healthy controls. Although it is tempting to hypothesize that activin A could have anti-inflammatory effects in patients with NAFLD, characterized by persistent inflammation as in patients with coronary heart disease, caution is need when interpreting the effect of activin A, not only on inflammation, but also on, for example, apoptosis. These effects are strongly dependent on cell type and not least the cell milieu, that is, affection on the cell from extrinsic and intrinsic factors.
V. Conclusion and Future Perspectives NAFLD is a major and steadily increasing worldwide cause of abnormal liver function, that ultimately may progress to NASH and end-stage liver disease. The pathogenesis of NAFLD development and progression is only partly understood, but involves IR, oxidative stress, apoptosis, fibrosis, and inflammation. Elucidation of central mediators of these processes is a necessary for designing therapy that targets the spectrum of disease that NAFLD represent. We have recently suggested that the TGF-b superfamily activin A may play a role in NAFLD pathogenesis based on both our findings of increased circulating levels of activin A and potentially increased hepatic activin A
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activity in NAFLD patients, and others and our own experimental observations. The role of activin A in NAFLD seems to be complex, involving both potentially beneficial effects on metabolism and detrimental effects such as hepatocyte apoptosis, fibrosis, and inflammation. Thus, based on the “twohit” hypothesis of NAFLD pathogenesis, it may seem that activin A may have beneficial effects on the “first hits” IR and fat accumulation (Fig. 15.2). In contrast, activin A may very well represent a mediator of the “second hits” that lead to progressive NAFLD, development of NASH, cirrhosis, and end-stage liver disease.
REFERENCES Adams, L. A., Sanderson, S., Lindor, K. D., and Angulo, P. (2005). The histological course of nonalcoholic fatty liver disease: A longitudinal study of 103 patients with sequential liver biopsies. J. Hepatol. 42(1), 132–138. Angulo, P. (2002). Nonalcoholic fatty liver disease. N. Engl. J. Med. 346(16), 1221–1231. Bataller, R., and Brenner, D. A. (2005). Liver fibrosis. J. Clin. Invest. 115(2), 209–218. Beier, K., Volkl, A., and Fahimi, H. D. (1992). Suppression of peroxisomal lipid betaoxidation enzymes of TNF-alpha. FEBS Lett. 310(3), 273–276. Bellentani, S., Saccoccio, G., Masutti, F., Croce, L. S., Brandi, G., Sasso, F., Cristanini, G., and Tiribelli, C. (2000). Prevalence of and risk factors for hepatic steatosis in Northern Italy. Ann. Intern. Med. 132(2), 112–117. Bjornsson, E., and Angulo, P. (2007). Non-alcoholic fatty liver disease. Scand. J. Gastroenterol. 42(9), 1023–1030. Browning, J. D., and Horton, J. D. (2004). Molecular mediators of hepatic steatosis and liver injury. J. Clin. Invest. 114(2), 147–152. Browning, J. D., Szczepaniak, L. S., Dobbins, R., Nuremberg, P., Horton, J. D., Cohen, J. C., Grundy, S. M., and Hobbs, H. H. (2004). Prevalence of hepatic steatosis in an urban population in the United States: Impact of ethnicity. Hepatology 40(6), 1387–1395. Bugianesi, E., Leone, N., Vanni, E., Marchesini, G., Brunello, F., Carucci, P., Musso, A., De Paolis, P., Capussotti, L., Salizzoni, M., et al. (2002). Expanding the natural history of nonalcoholic steatohepatitis: From cryptogenic cirrhosis to hepatocellular carcinoma. Gastroenterology 123(1), 134–140. Bugianesi, E., Gastaldelli, A., Vanni, E., Gambino, R., Cassader, M., Baldi, S., Ponti, V., Pagano, G., Ferrannini, E., and Rizzetto, M. (2005). Insulin resistance in non-diabetic patients with non-alcoholic fatty liver disease: Sites and mechanisms. Diabetologia 48(4), 634–642. Canbay, A., Taimr, P., Torok, N., Higuchi, H., Friedman, S., and Gores, G. J. (2003). Apoptotic body engulfment by a human stellate cell line is profibrogenic. Lab. Invest. 83 (5), 655–663. Cerami, A., and Beutler, B. (1988). The role of cachectin/TNF in endotoxic shock and cachexia. Immunol. Today 9(1), 28–31. Chen, Y. G., Lui, H. M., Lin, S. L., Lee, J. M., and Ying, S. Y. (2002). Regulation of cell proliferation, apoptosis, and carcinogenesis by activin. Exp. Biol. Med. (Maywood) 227(2), 75–87. Cortez-Pinto, H., de Moura, M. C., and Day, C. P. (2006). Non-alcoholic steatohepatitis: From cell biology to clinical practice. J. Hepatol. 44(1), 197–208.
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Dam-Larsen, S., Franzmann, M., Andersen, I. B., Christoffersen, P., Jensen, L. B., Sorensen, T. I., Becker, U., and Bendtsen, F. (2004). Long term prognosis of fatty liver: Risk of chronic liver disease and death. Gut 53(5), 750–755. Dandona, P., Aljada, A., Chaudhuri, A., Mohanty, P., and Garg, R. (2005). Metabolic syndrome: A comprehensive perspective based on interactions between obesity, diabetes, and inflammation. Circulation 111(11), 1448–1454. Day, C. (2007). Metabolic syndrome, or what you will: Definitions and epidemiology. Diab. Vasc. Dis. Res. 4(1), 32–38. Day, C. P., and James, O. F. (1998). Steatohepatitis: A tale of two “hits”? Gastroenterology 114(4), 842–845. De Bleser, P. J., Niki, T., Xu, G., Rogiers, V., and Geerts, A. (1997). Localization and cellular sources of activins in normal and fibrotic rat liver. Hepatology 26(4), 905–912. de Caestecker, M. (2004). The transforming growth factor-beta superfamily of receptors. Cytokine Growth Factor Rev. 15(1), 1–11. Deli, A., Kreidl, E., Santifaller, S., Trotter, B., Seir, K., Berger, W., Schulte-Hermann, R., Rodgarkia-Dara, C., and Grusch, M. (2008). Activins and activin antagonists in hepatocellular carcinoma. World J. Gastroenterol. 14(11), 1699–1709. den Boer, M. A., Voshol, P. J., Schroder-van der Elst, J. P., Korsheninnikova, E., Ouwens, D. M., Kuipers, F., Havekes, L. M., and Romijn, J. A. (2006). Endogenous interleukin-10 protects against hepatic steatosis but does not improve insulin sensitivity during high-fat feeding in mice. Endocrinology 147(10), 4553–4558. Donati, G., Stagni, B., Piscaglia, F., Venturoli, N., Morselli-Labate, A. M., Rasciti, L., and Bolondi, L. (2004). Increased prevalence of fatty liver in arterial hypertensive patients with normal liver enzymes: Role of insulin resistance. Gut 53(7), 1020–1023. Donnelly, K. L., Smith, C. I., Schwarzenberg, S. J., Jessurun, J., Boldt, M. D., and Parks, E. J. (2005). Sources of fatty acids stored in liver and secreted via lipoproteins in patients with nonalcoholic fatty liver disease. J. Clin. Invest. 115(5), 1343–1351. Ekstedt, M., Franzen, L. E., Mathiesen, U. L., Thorelius, L., Holmqvist, M., Bodemar, G., and Kechagias, S. (2006). Long-term follow-up of patients with NAFLD and elevated liver enzymes. Hepatology 44(4), 865–873. Fadok, V. A., Bratton, D. L., Konowal, A., Freed, P. W., Westcott, J. Y., and Henson, P. M. (1998). Macrophages that have ingested apoptotic cells in vitro inhibit proinflammatory cytokine production through autocrine/paracrine mechanisms involving TGF-beta, PGE2, and PAF. J. Clin. Invest. 101(4), 890–898. Fassio, E., Alvarez, E., Dominguez, N., Landeira, G., and Longo, C. (2004). Natural history of nonalcoholic steatohepatitis: A longitudinal study of repeat liver biopsies. Hepatology 40(4), 820–826. Festa, A., D0 Agostino, R., Jr., Howard, G., Mykkanen, L., Tracy, R. P., and Haffner, S. M. (2000). Chronic subclinical inflammation as part of the insulin resistance syndrome: The Insulin Resistance Atherosclerosis Study (IRAS). Circulation 102(1), 42–47. Fromenty, B., Robin, M. A., Igoudjil, A., Mansouri, A., and Pessayre, D. (2004). The ins and outs of mitochondrial dysfunction in NASH. Diabetes Metab. 30(2), 121–138. Furukawa, M., Nobusawa, R., Shibata, H., Eto, Y., and Kojima, I. (1995). Initiation of insulin secretion in glucose-free medium by activin A. Mol. Cell. Endocrinol. 113(1), 83–87. Gold, E. J., Francis, R. J., Zimmermann, A., Mellor, S. L., Cranfield, M., Risbridger, G. P., Groome, N. P., Wheatley, A. M., and Fleming, J. S. (2003). Changes in activin and activin receptor subunit expression in rat liver during the development of CCl4-induced cirrhosis. Mol. Cell. Endocrinol. 201(1–2), 143–153. Gressner, A. M., Lahme, B., Mannherz, H. G., and Polzar, B. (1997). TGF-beta-mediated hepatocellular apoptosis by rat and human hepatoma cells and primary rat hepatocytes. J. Hepatol. 26(5), 1079–1092.
340
Arne Yndestad et al.
Gressner, O. A., Lahme, B., Siluschek, M., Rehbein, K., Weiskirchen, R., and Gressner, A. M. (2008). Intracrine signalling of activin A in hepatocytes upregulates connective tissue growth factor (CTGF/CCN2) expression. Liver Int. 28(9), 1207–1216. Grusch, M., Drucker, C., Peter-Vorosmarty, B., Erlach, N., Lackner, A., Losert, A., Macheiner, D., Schneider, W. J., Hermann, M., Groome, N. P., et al. (2006). Deregulation of the activin/follistatin system in hepatocarcinogenesis. J. Hepatol. 45(5), 673–680. Gupte, P., Amarapurkar, D., Agal, S., Baijal, R., Kulshrestha, P., Pramanik, S., Patel, N., Madan, A., Amarapurkar, A., and Hafeezunnisa, A. (2004). Non-alcoholic steatohepatitis in type 2 diabetes mellitus. J. Gastroenterol. Hepatol. 19(8), 854–858. Han, Y. P. (2006). Matrix metalloproteinases, the pros and cons, in liver fibrosis. J. Gastroenterol. Hepatol. 21(Suppl. 3), S88–S91. Harrison, S. A., Torgerson, S., and Hayashi, P. H. (2003). The natural history of nonalcoholic fatty liver disease: A clinical histopathological study. Am. J. Gastroenterol. 98(9), 2042–2047. Haukeland, J. W., Damas, J. K., Konopski, Z., Loberg, E. M., Haaland, T., Goverud, I., Torjesen, P. A., Birkeland, K., Bjoro, K., and Aukrust, P. (2006). Systemic inflammation in nonalcoholic fatty liver disease is characterized by elevated levels of CCL2. J. Hepatol. 44(6), 1167–1174. Horton, J. D., Goldstein, J. L., and Brown, M. S. (2002). SREBPs: Activators of the complete program of cholesterol and fatty acid synthesis in the liver. J. Clin. Invest. 109 (9), 1125–1131. Hube, F., and Hauner, H. (1999). The role of TNF-alpha in human adipose tissue: Prevention of weight gain at the expense of insulin resistance? Horm. Metab. Res. 31 (12), 626–631. Hubner, G., Alzheimer, C., and Werner, S. (1999). Activin: A novel player in tissue repair processes. Histol. Histopathol. 14(1), 295–304. Hughes, R. D., and Evans, L. W. (2003). Activin A and follistatin in acute liver failure. Eur. J. Gastroenterol. Hepatol. 15(2), 127–131. Iredale, J. P. (2007). Models of liver fibrosis: Exploring the dynamic nature of inflammation and repair in a solid organ. J. Clin. Invest. 117(3), 539–548. Jones, K. L., de Kretser, D. M., Patella, S., and Phillips, D. J. (2004). Activin A and follistatin in systemic inflammation. Mol. Cell. Endocrinol. 225(1–2), 119–125. Jou, J., Choi, S. S., and Diehl, A. M. (2008). Mechanisms of disease progression in nonalcoholic fatty liver disease. Semin. Liver Dis. 28(4), 370–379. Kanda, H., Tateya, S., Tamori, Y., Kotani, K., Hiasa, K., Kitazawa, R., Kitazawa, S., Miyachi, H., Maeda, S., Egashira, K., et al. (2006). MCP-1 contributes to macrophage infiltration into adipose tissue, insulin resistance, and hepatic steatosis in obesity. J. Clin. Invest. 116(6), 1494–1505. Kerner, J., and Hoppel, C. (2000). Fatty acid import into mitochondria. Biochim. Biophys. Acta 1486(1), 1–17. Larter, C. Z., Yeh, M. M., Haigh, W. G., Williams, J., Brown, S., Bell-Anderson, K. S., Lee, S. P., and Farrell, G. C. (2008). Hepatic free fatty acids accumulate in experimental steatohepatitis: Role of adaptive pathways. J. Hepatol. 48(4), 638–647. Lee, J. Y., Kim, K. M., Lee, S. G., Yu, E., Lim, Y. S., Lee, H. C., Chung, Y. H., Lee, Y. S., and Suh, D. J. (2007). Prevalence and risk factors of non-alcoholic fatty liver disease in potential living liver donors in Korea: A review of 589 consecutive liver biopsies in a single center. J. Hepatol. 47(2), 239–244. Lewis, J. R., and Mohanty, S. R. (2010). Nonalcoholic fatty liver disease: A review and update. Dig. Dis. Sci. 55(3), 560–578. Li, L., Yi, Z., Seno, M., and Kojima, I. (2004). Activin A and betacellulin: Effect on regeneration of pancreatic beta-cells in neonatal streptozotocin-treated rats. Diabetes 53 (3), 608–615.
Activin A in Nonalcoholic Fatty Liver Disease
341
Lin, S. D., Kawakami, T., Ushio, A., Sato, A., Sato, S., Iwai, M., Endo, R., Takikawa, Y., and Suzuki, K. (2006). Ratio of circulating follistatin and activin A reflects the severity of acute liver injury and prognosis in patients with acute liver failure. J. Gastroenterol. Hepatol. 21(2), 374–380. Lind, P. (2004). Interdependence of hepatic lipid and glucose metabolism: Novel pharmacological targets for diabetes. Curr. Opin. Investig. Drugs 5(4), 395–401. Ling, N., Ying, S. Y., Ueno, N., Shimasaki, S., Esch, F., Hotta, M., and Guillemin, R. (1986). Pituitary FSH is released by a heterodimer of the beta-subunits from the two forms of inhibin. Nature 321(6072), 779–782. Loria, P., Lonardo, A., and Carulli, N. (2005). Should nonalcoholic fatty liver disease be renamed? Dig. Dis. 23(1), 72–82. Ludwig, J., Viggiano, T. R., McGill, D. B., and Oh, B. J. (1980). Nonalcoholic steatohepatitis: Mayo Clinic experiences with a hitherto unnamed disease. Mayo Clin. Proc. 55(7), 434–438. Machado, M., Marques-Vidal, P., and Cortez-Pinto, H. (2006). Hepatic histology in obese patients undergoing bariatric surgery. J. Hepatol. 45(4), 600–606. Marchesini, G., Bugianesi, E., Forlani, G., Cerrelli, F., Lenzi, M., Manini, R., Natale, S., Vanni, E., Villanova, N., Melchionda, N., et al. (2003). Nonalcoholic fatty liver, steatohepatitis, and the metabolic syndrome. Hepatology 37(4), 917–923. Mukherjee, A., Sidis, Y., Mahan, A., Raher, M. J., Xia, Y., Rosen, E. D., Bloch, K. D., Thomas, M. K., and Schneyer, A. L. (2007). FSTL3 deletion reveals roles for TGF-beta family ligands in glucose and fat homeostasis in adults. Proc. Natl. Acad. Sci. USA 104(4), 1348–1353. Pagano, G., Pacini, G., Musso, G., Gambino, R., Mecca, F., Depetris, N., Cassader, M., David, E., Cavallo-Perin, P., and Rizzetto, M. (2002). Nonalcoholic steatohepatitis, insulin resistance, and metabolic syndrome: Further evidence for an etiologic association. Hepatology 35(2), 367–372. Pangas, S. A., and Woodruff, T. K. (2000). Activin signal transduction pathways. Trends Endocrinol. Metab. 11(8), 309–314. Patella, S., Phillips, D. J., de Kretser, D. M., Evans, L. W., Groome, N. P., and Sievert, W. (2001). Characterization of serum activin-A and follistatin and their relation to virological and histological determinants in chronic viral hepatitis. J. Hepatol. 34(4), 576–583. Patella, S., Phillips, D. J., Tchongue, J., de Kretser, D. M., and Sievert, W. (2006). Follistatin attenuates early liver fibrosis: Effects on hepatic stellate cell activation and hepatocyte apoptosis. Am. J. Physiol. Gastrointest. Liver Physiol. 290(1), G137–G144. Piek, E., Heldin, C. H., and Ten Dijke, P. (1999). Specificity, diversity, and regulation in TGF-beta superfamily signaling. FASEB J. 13(15), 2105–2124. Rodgarkia-Dara, C., Vejda, S., Erlach, N., Losert, A., Bursch, W., Berger, W., SchulteHermann, R., and Grusch, M. (2006). The activin axis in liver biology and disease. Mutat. Res. 613(2–3), 123–137. Russell, C. E., Hedger, M. P., Brauman, J. N., de Kretser, D. M., and Phillips, D. J. (1999). Activin A regulates growth and acute phase proteins in the human liver cell line, HepG2. Mol. Cell. Endocrinol. 148(1–2), 129–136. Sanyal, A. J., Campbell-Sargent, C., Mirshahi, F., Rizzo, W. B., Contos, M. J., Sterling, R. K., Luketic, V. A., Shiffman, M. L., and Clore, J. N. (2001). Nonalcoholic steatohepatitis: Association of insulin resistance and mitochondrial abnormalities. Gastroenterology 120(5), 1183–1192. Schwall, R. H., Robbins, K., Jardieu, P., Chang, L., Lai, C., and Terrell, T. G. (1993). Activin induces cell death in hepatocytes in vivo and in vitro. Hepatology 18(2), 347–356. Seki, S., Kitada, T., Yamada, T., Sakaguchi, H., Nakatani, K., and Wakasa, K. (2002). In situ detection of lipid peroxidation and oxidative DNA damage in non-alcoholic fatty liver diseases. J. Hepatol. 37(1), 56–62.
342
Arne Yndestad et al.
Seppala-Lindroos, A., Vehkavaara, S., Hakkinen, A. M., Goto, T., Westerbacka, J., Sovijarvi, A., Halavaara, J., and Yki-Jarvinen, H. (2002). Fat accumulation in the liver is associated with defects in insulin suppression of glucose production and serum free fatty acids independent of obesity in normal men. J. Clin. Endocrinol. Metab. 87(7), 3023–3028. Shimomura, I., Matsuda, M., Hammer, R. E., Bashmakov, Y., Brown, M. S., and Goldstein, J. L. (2000). Decreased IRS-2 and increased SREBP-1c lead to mixed insulin resistance and sensitivity in livers of lipodystrophic and ob/ob mice. Mol. Cell 6(1), 77–86. Smith, C., Yndestad, A., Halvorsen, B., Ueland, T., Waehre, T., Otterdal, K., Scholz, H., Endresen, K., Gullestad, L., Froland, S. S., et al. (2004). Potential anti-inflammatory role of activin A in acute coronary syndromes. J. Am. Coll. Cardiol. 44(2), 369–375. Tanaka, N., Sano, K., Horiuchi, A., Tanaka, E., Kiyosawa, K., and Aoyama, T. (2008). Highly purified eicosapentaenoic acid treatment improves nonalcoholic steatohepatitis. J. Clin. Gastroenterol. 42(4), 413–418. Teli, M. R., James, O. F., Burt, A. D., Bennett, M. K., and Day, C. P. (1995). The natural history of nonalcoholic fatty liver: A follow-up study. Hepatology 22(6), 1714–1719. Tilg, H., and Moschen, A. R. (2008). Insulin resistance, inflammation, and non-alcoholic fatty liver disease. Trends Endocrinol. Metab. 19(10), 371–379. Ueda, Y. (2000). Activin A increases Pax4 gene expression in pancreatic beta cell lines. FEBS Lett. 480(2–3), 101–105. Unger, R. H., and Orci, L. (2002). Lipoapoptosis: Its mechanism and its diseases. Biochim. Biophys. Acta 1585(2–3), 202–212. Uysal, K. T., Wiesbrock, S. M., Marino, M. W., and Hotamisligil, G. S. (1997). Protection from obesity-induced insulin resistance in mice lacking TNF-alpha function. Nature 389 (6651), 610–614. Wang, M., Liu, A., Garcia, F. U., Rhim, J. S., and Stearns, M. E. (1999). Growth of HPV18 immortalized human prostatic intraepithelial neoplasia cell lines. Influence of IL-10, follistatin, activin-A, and DHT. Int. J. Oncol. 14(6), 1185–1195. Wanless, I. R., and Lentz, J. S. (1990). Fatty liver hepatitis (steatohepatitis) and obesity: An autopsy study with analysis of risk factors. Hepatology 12(5), 1106–1110. Weisberg, S. P., Hunter, D., Huber, R., Lemieux, J., Slaymaker, S., Vaddi, K., Charo, I., Leibel, R. L., and Ferrante, A. W., Jr. (2006). CCR2 modulates inflammatory and metabolic effects of high-fat feeding. J. Clin. Invest. 116(1), 115–124. Wellen, K. E., and Hotamisligil, G. S. (2005). Inflammation, stress, and diabetes. J. Clin. Invest. 115(5), 1111–1119. Werner, S., and Alzheimer, C. (2006). Roles of activin in tissue repair, fibrosis, and inflammatory disease. Cytokine Growth Factor Rev. 17(3), 157–171. Yamaoka, T., Idehara, C., Yano, M., Matsushita, T., Yamada, T., Ii, S., Moritani, M., Hata, J., Sugino, H., Noji, S., et al. (1998). Hypoplasia of pancreatic islets in transgenic mice expressing activin receptor mutants. J. Clin. Invest. 102(2), 294–301. Yeh, M. M., and Brunt, E. M. (2007). Pathology of nonalcoholic fatty liver disease. Am. J. Clin. Pathol. 128(5), 837–847. Yndestad, A., Ueland, T., ie, E., Florholmen, G., Halvorsen, B., Attramadal, H., Simonsen, S., Fr land, S. S., Gullestad, L., Christensen, G., et al. (2004). Elevated levels of activin A in heart failure: Potential role in myocardial remodeling. Circulation 109(11), 1379–1385. Yndestad, A., Haukeland, J. W., Dahl, T. B., Bjoro, K., Gladhaug, I. P., Berge, C., Damas, J. K., Haaland, T., Loberg, E. M., Linnestad, P., et al. (2009). A complex role of activin A in non-alcoholic fatty liver disease. Am. J. Gastroenterol. 104(9), 2196–2205. Yuen, M. F., Norris, S., Evans, L. W., Langley, P. G., and Hughes, R. D. (2002). Transforming growth factor-beta 1, activin and follistatin in patients with hepatocellular carcinoma and patients with alcoholic cirrhosis. Scand. J. Gastroenterol. 37(2), 233–238. Zelman, S. (1952). The liver in obesity. AMA Arch. Intern. Med. 90(2), 141–156.
Index
A Activin activin A gene, 260–261 bA-subunit, 260 bB-subunit, 260–261 bD subunit, 261 bioassays, 258–259 CNS (see also Brain, activin) depression and anxiety-related behavior, 190–193 late-phase LTP, 195–196 postnatal neurogenesis, 193–195 reconsolidation and extinction, 197–200 spine formation, 189–190 definition, 5 follistatin binding affinity, 263 FS288, 263 FS315, 263–264 FSRP/FSTL3, 264 glycosylation, 264–265 introns and exons, 262 gene disruption studies, 6 glucose metabolism (see Glucose metabolism, activins) HEK293 and HT22 cells, 6–7 immune cell migration dendritic cells, 141 mast cells, 139–140 monocytes, 140–141 immunohistochemistry, 258 isoforms, 5–9 molecular mechanism Smads-dependent cell migration, 132–133 Smads-independent cell migration, 133–134 nomenclature, 260 pancreatic b-cell line MIN6 cells, 6 receptors and activin signaling BAMBI, 262 betaglycan, 262 MAP kinases, 261–262 SMAD2/3 signaling, 263 signaling cascade, postsynaptic region, 186–187 structure, 186 synthesis and signaling, 258, 259 tumor cell migration breast cancer, 135–137
colon cancer, 138 prostate cancer, 134–135 Activin A bioassays, 258–259 cachexia, 270–271 continuous low-level activation, 63–65 follistatin and FSTL3, 330–331 production site and measurement issues, 164–165 gene, 260 immunohistochemistry, 258 immunoregulation, 279 asthma, 278–279 B cell survival, 275 dendritic and natural killer cells, 278 IL6 production, 277 MHC class II proteins, 275 monocyte/macrophage production, 276 multipotential progenitor cell development, 275 myeloid dendritic cells, 276 T cell growth and development, 275 inflammation and fibrotic response, follistatin bC-subunit, 274 exogenous follistatin, 273–274 monocyte/macrophage lineage cell types, 271–273 stimulating factors, 273 inflammation and immunity acute and chronic inflammatory diseases, 265 cell types, activin production, 266 cytokines, 269 interleukin-1, 267–268 LPS injection, 266 mRNA and protein elevation, 265–266 NF-kB signaling, 268 oxidative stress, 269 phorbol ester responsive elements, 268–269 septicemia and pulmonary hypertension, 265 toll-like receptor signaling pathway, 267–268 intracrine signaling, 71–73 NAFLD apoptosis and inhibition, hepatocyte growth, 331–332
343
344 Activin A (cont.) metabolic syndrome (see Metabolic syndrome, NAFLD) two-hit hypothesis, 332, 333 nomenclature, 258 properties, 329 signal transduction and regulation, 329–330 Smad2/3 pathway, 331 synthesis, 258 Activin B inflammation and immunity, 269–270 Activin receptor and BMP coreceptors, 110–111 type II receptors, 110 type I receptors, 109–110 function activin membrane-bound inhibitor, 85 BMP, 85 Cripto, 85–87 Dapper2, 89 Dok-PTB containing protein, 88–89 proteins 1 and 2, 87–88 regulatory subunits of PP2A, 87 and their receptors, 107–108 Activin receptor interacting proteins (ARIPs), 87–88 Activin receptor-like kinase (ALK), 109–110, 131, 219 activin/nodal and BMP signaling, 209–210 activins A and B signal transduction, 303 ActRIIA and ActRIIB, 30 ALK4 and 7, 6–7 BMP-2, 115 BMP signaling, 4 BMP2/4/7 signaling, 68, 69 breast cancer, 137 Cripto, 86 CTGF expression, 67 Dapper2, 89 dendritic cells, 141 IL6 production, 277 inhibitory Smads, 89 insulin secretion, 220–221 myostatin signal transduction pathway, 223, 274 nodal and activin pathway, 47 nodal signaling, 9 pancreatic endocrine cells, 7–8 prostate cancer cell, 132 SB-431542 and GW788388, 201 Smad2 and 3 phosphorylation, 14, 15 Smads-dependent cell migration, 132 stimulation, intracellular TGF-b signaling, 68 structure, 4 white adipose tissue, 224 Activin receptor type IB (ActRIB) FKBP12 and dorsomorphin, 31–32
Index
TGF-b RI kinase domain structure, 31 Activin receptor type IIB (ActRIIB) active site of, 33–35 catalytic domain of, 32–33 vs. type IB domain structures, 35–36 Anti-Mullerian hormone (AMH) follicular phase, 307 FSH human studies, 310 inhibin B, 307–308 inhibins and LH, 310 ovulatory cycles, 307 Anxiety-related behavior. See Depression and anxiety-related behavior Apoptosis, 328, 335 B B cells, activin A, 239–240 Betaglycan inhibins A and B ActRII complex, 314–316 b-subunits, 313 vs. diglycosylated (34-kDa) inhibins A and B, 312–313 epitope, 313–314 inhibin B specific accessory protein, 314, 315 mRNA expression, 169 Bone marrow-derived mast cells (BMMCs) chemoattractant, 245 MCP-6 and MCP-7, 244 regulatory effects, 244 Smad3, 246 Bone morphogenic proteins (BMPs), 2 ActRIIBECD, 111 ALK1, 2, 3, and 6, 4 binding affinity, 114–115 BMP-6, 154 chimeras, 116, 117 coreceptors, 110–111 GDF3, 49 inhibitory Smads, 89 ligand flexibility, 112–113 mouse and human ESCs, 44–45 negative regulation, 85 sequence alignment, 114 Ski and SnoN, 94 stem cell pluripotency, 51 structural studies architecture of complex structures, 111–112 available complex structures, 111 binding epitopes, 113–115 ligand flexibility, 112–113 receptor-receptor interaction, 112 type II receptors, 110 type I receptors, 109–110 Brain, activin activin type II receptor, 189
345
Index
adult neurogenesis bromodeoxyuridine, 193 electroconvulsive therapy, 195 excitatory and inhibitory postsynaptic potential, 194–195 follistatin and GDF11, 194 FSM/ACM-double transgenic mice, 194 hippocampal culture, 194 physiological roles, 193 depression and anxiety-related behavior antidepressant drugs, 193 dominant-negative ActRIB, 190–191, 193 FSM and ACM, 190, 192 GKAP, 189 glutamate, 189 late-phase LTP, 195–196 memory reconsolidation and extinction fear conditioning test, 197–200 posttraumatic stress disorder, 197 three-week memory testing, 197 NMDA receptor, 189 postsynaptic density 95 (PSD95), 189 spine formation plasticity, 189 presynaptic contacts, 190 Smad pathway, 190 spine morphology, 188 Breast cancer, 135–137, 143 Bromodeoxyuridine, 193 Brown adipose tissue, 225 C cAMP response element (CRE), 12 Canonical Wnt signaling, 210–212 Cell migration immune cells dendritic cells, 141 mast cells, 139–140 monocytes, 140–141 molecular mechanism of activin signaling Smads-dependent cell migration, 132–133 Smads-independent cell migration, 133–134 tumors breast cancer, 135–137 colon cancer, 138 prostate cancer, 134–135 Chaperones, 164 Chimeras Activin-like bioactivity, 116–118 ActRII binding properties, 116 antagonism, 118–119 BMP-like bioactivity, 119–121 design of, 115–116 Colon cancer, 138, 143 Connective tissue growth factor (CTGF), 60–61 Contextual fear-conditioning test, 197, 199 ABItTA mice, 200
FBItTA mice, 197, 199 Cripto, 85–87, 108 D Dapper 2, 89 Dendritic cells (DCs), 141, 143 Depression and anxiety-related behavior antidepressant drugs, 193 dominant-negative ActRIB, 190–191, 193 FSM and ACM, 190, 192 Diglycosylated (34-kDa) inhibins A and B, 312–313 Dok-PTB containing protein (Dpcp), 88–89 E Ectodermin/TIF1g, 91–92 Electroconvulsive therapy (ECT), 195 Embryonic stem cells (ESCs) derivation of, 43–44 induced-pluripotent stem cells, 45–46 locations of pluripotent cells, 41–42 mouse and human, 44–45 Endoderm formation activin/nodal and BMP signaling, 209–210 differentiation, 208 Wnt signaling pathway canonical, 210–212 hepatocyte-like cells (see Hepatocyte-like cells (HLCs)) noncanonical, 211–212 Endoglin, transmembrane protein, 111 Epigenetic regulation, inhibin histone modifications, 154–155 hypermethylation, 154 Epithelial-mesenchymal transition (EMT), 132–133 Erbin protein, 93–94 Evi-1, zinc finger-containing transcriptional factor, 96 Exogenous FSH stimulation, 305 Extracellular antagonists follistatin-related gene, 83–85 inhibin, 82–83 Extragonadal expression, inhibin female reproduction, 158–159 target receptors expression, 158 tissue and cellular localization, 159 F Fear conditioning test, 197–200 Fibrodysplasia ossificans progressiva (FOP), 31 Follicle-stimulating hormones (FSHs), 30 AMH, 309–310 human menstrual cycle follicular phase, 305, 307 luteal phase, 305
346
Index
Follicle-stimulating hormones (FSHs) (cont.) ovarian and pituitary hormones, changing patterns, 310–311 inhibin A and, 309 inhibin B and, 311–313 inhibins A and B bioactivity ALK4 and ALK7 type I receptors, 316–317 antagonism, 314–316 betaglycan, 312–314 glycosylation, 314 mono and diglycosylated forms, 316 posttranslational modifications, 316 LH, 310–311 male, inhibins and regulation, 311–312 Follicular phase, human menstrual cycle, 305, 307 Follistatin binding affinity, 263–264 FS288, 263 FS315, 263–264 inflammation and fibrotic response bC-subunit, 274 exogenous follistatin, 273–274 monocyte/macrophage lineage cell types, 271–273 stimulating factors, 273 production site and measurement issues, 264–265 structure, 262–263 Follistatin-related gene (FLRG), 83–85 G GATA factor, 151, 152 Glucose metabolism, activins, 227, 334 activins A, 226 activins C and E, 226–227 adipose tissues brown, 225 inflammation, macrophages, 225–226 white, 223–224 blood glucose level, 220 carbohydrates consumption, 220 hypertrophy, Akt/mTOR pathway activation, 228 liver activin A, 221–222 activins C and E, 222 gluconeogenesis, 221–222 glycogenolysis, 221 mitogen-stimulated growth inhibition, 221 obesity, 220 pancreas adult ActRIIA-or ActRIIB-null mice, 219–220 ALK7, insulin gene transcription, 221 ES cell line, 220 glucagon expression, 221 insulin secretion, 220–221 regeneration model, 220
phosphatidylinositol-3 kinase (PI3K) pathways, 227 skeletal muscle ActRIIB/Fc fusion protein, 223 C2C12 myoblast cells, 222–223 muscle mass preservation, 222 myostatin, 223 tibialis anterior muscle, 222 Gonadal steroids, 300 Gonadotrophins, inhibin cAMP levels, 151–153 CREB phosphorylation, 151–152 FSH production, 153 gonadotrophin-releasing hormone, 153 Granulosa cell tumors, 169 Guanylate kinase domain-associated protein (GKAP), 189 H Hepatic fibrosis matrix metalloproteinases, EMC, 336 paracrine and autocrine signaling, 336 tissue homeostasis, 335–336 Hepatocyte-like cells (HLCs) bioartificial liver device, 213 human drug toxicity, 213 human liver function model, 212–213 isolation, 212 Histone modifications, inhibin, 154–155 Human embryonic stem cells (hESCs), 44–45 Humoral immune responses, activin A B cells, 239–240 macrophages cytokines and chemokines, 240 iNOS and arginase, 242, 243 monocyte-derived dendritic cells, 240–241 peripheral blood myeloid dendritic cells, 240–241 TGF-b gene transcripts, 241 type IV collagenase production, 241–242 mast cells modulation, 244–246 upregulation, 242–244 TGF-b, 236–237 Th2 cells activin bA promoter, transactivation, 238 CD4þ CD4-T cells, 237–238 expression, T helper cells, 237 Hypermethylation, inhibin, 154 Hypothalamic pituitary gonadal axis (HPG), 158 I Immune cell migration dendritic cells, 141 mast cells, 139–140 monocytes, 140–141 Induced-pluripotent stem cells (iPSCs), 45–46
347
Index
Inflammation, 336–337 activins bB-subunit, 269–270 interleukin-1 (IL1), 267–270 LPS injection, 266 mRNA expression and protein localization, 265–266 toll-like receptor, 267–270 follistatin, 270 Inhibin, 82–83, 302 biological actions, 150 chaperones, 164 circulating forms B forms, men, 170 ovarian cancer, 166, 169 women, 166–168 expression and regulation bB-subunit, 155–154 BMP-6, 154 epigenetic regulation, 154–155 GATA factor, 151, 152 gonadotrophins, 151–153 human inhibin, 151 inhibin synthesis, 151 posttranscriptional regulation, 155, 170–171 Smad-binding element, 153 TATA boxes, 151, 152 5’ untranslated regions (UTRs), 151, 152 FSH suppression (see FSH suppression, inhibin) human tissues adult, 155, 156 extragonadal expression, 158–159 females, 154, 157 HPG axis, 158 males, 157–158 inhibin A, molecular mass, 167–168 mechanism of action activins A and B signal transduction, 303 antagonism, 303–304 FSHb subunit synthesis, 304 LHb subunit synthesis, 305 posttranslational modifications, 163–164 prodomains, sequence alignment, 160, 162 protein structure, 159–162 proteolytic processing, 162–163 regulation, 150 structure glycosylation sites, 302 heterodimers, 301–302 homodimers, 302 precursor inhibin molecule, 300 TGFb isoform, 302–303 subunits, 150 synthesis and secretion, 160, 161 Inhibitory Smads (I-Smads), 47, 89–90 Insulin gene regulation
cAMP response element (CRE), 12 C element, 12–13 E element, 13–14 A element, 10 GG element, 11–12 Smad-binding element (SBE), 14 Intracrine signaling mechanism Activin A, 72–73 aspects of, 65–66 TGF-b continuous low-level activation, 63–65 external and internal inhibitors, 65 inhibitory pathway, 70–72 intracellular activation, 61–63 stimulatory pathway, 66–70 Ionotropic glutamate receptors, 189 L Latent TGF-b binding proteins (LTBPs), 164 Late-phase LTP, 195–196 Liver activin A gluconeogenesis, 221–222 glycogenolysis, 221 mitogen-stimulated growth inhibition, 221 activins C and E, 222 M Macrophages, activin A cytokines and chemokines, 240 iNOS and arginase, 242, 243 monocyte-derived dendritic cells, 240–241 peripheral blood myeloid dendritic cells, 240–241 TGF-b gene transcripts, 241 type IV collagenase production, 241–242 Mast cells (MCs), 139–140, 143 BMMC maturation chemoattractant, 245 MCP-6 and MCP-7, 244 regulatory effects, 244 Smad3, 246 upregulation antigen receptors, cytosolic Ca2þ levels, 243 high-affinity IgE receptor, 242–243 mast cell maturation, 243 Matrix metalloproteinases (MMPs), 241, 336 Metabolic syndrome, NAFLD activin A activin A/follistatin ratio, 332 apoptosis, 335 glucose metabolism, 334 hepatic FA metabolism, 333–334 hepatic fibrosis, 335–336 inflammation, 336–337 steatosis, 332–333
348
Index
Metabolic syndrome, NAFLD (cont.) type 2 diabetes mellitus, 326–327 Microphthalmia-associated transcription factor (MITF), 244 MicroRNAs (miRNAs), 155 Miscarriage, 159 Monocyte-derived dendritic cells, 240–241 Monocytes, 140–141, 143 Mouse embryonic fibroblasts (MEFs), 43 Mouse embryonic stem cells (mESCs), 44–45 N Negative regulation activin receptor function activin membrane-bound inhibitor, 85 BMP, 85 Cripto, 85–87 Dapper2, 89 Dok-PTB containing protein, 88–89 proteins 1 and 2, 87–88 regulatory subunits of PP2A, 87 gene transcription, 94–97 Neurogenesis, activin bromodeoxyuridine, 193 electroconvulsive therapy, 195 excitatory and inhibitory postsynaptic potential, 194–195 follistatin and GDF11, 194 FSM/ACM-double transgenic mice, 194 hippocampal culture, 194 physiological roles, 193 N-linked glycosylation, inhibin, 163 NMDA receptor, 189 Nodal signaling, 9 Nonalcoholic fatty liver disease (NAFLD) apoptosis, 328 diagnosis, 325 epidemiology, 326 fibrosis, 328 hepatic fat accumulation, 326–327 histology, 324 lipid and glucose metabolism, insulin, 327 metabolic syndrome, 326–327 natural history, 325 pathophysiology, 326 prevalence, 326 systemic inflammation, 328–329 two-hit model, 326, 340 Noncanonical Wnt signaling, 211–212 Nuclear factor of activated T cells (NFAT), 238 O Ovarian and pituitary hormones, changing patterns, 310–311 Ovarian cancer, 166, 169
P Pancreatic endocrine cells, 7–9 Pathogen-associated molecular patterns (PAMPs), 267 Peripheral blood mononuclear cells (PBMC), 337 Peripheral blood myeloid dendritic cells, 240–241 Placentation and pregnancy, inhibin, 158–159 Plasminogen activator inhibitor-1 (PAI-1), 224 Pluripotent cells. See Embryonic stem cells (ESCs) Postsynaptic density 95 (PSD95), 189 Posttranscriptional regulation, inhibin, 155 Posttranslational modifications, inhibin, 163–164 Preeclampsia, 159 Prostate cancer, 134–135, 143 R Receptor-activated SMADs (R-SMADs), 47, 60 S Serum response factor (SRF), 96 SMAD anchor for receptor activation (SARA), 210 Smad-binding element (SBE), 14, 153–154 Smad proteins, 47 cell migration Smads-dependent, 132–133 Smads-independent, 133–134 function ectodermin/TIF1g, 91–92 Erbin, 93–94 linker phosphorylation, 92–93 PPM1A, 91 transmembrane prostate androgen-induced RNA, 94 signaling pathway, 190 Smad2/3 signaling pathway, 331 Spine formation, activin, 189–190 Stem cell pluripotency Activin/Nodal/TGFb pathway, 46–48 TGFb signaling, 50–51 T TATA boxes, 151, 152 Th2 cells, activin A activin bA promoter, transactivation, 238 CD4þ CD4-T cells, 237–238 expression, T helper cells, 237 Transcription regulation cAMP response element (CRE), 12 C element, 12–13 E element, 13–14 A element, 10 GG element, 11–12 Smad-binding element (SBE), 14 Transforming growth factor-b (TGF-b)
349
Index
connective tissue growth factor, 60–61 continuous low-level activation, 63–65 external and internal inhibitors, 65 intracellular activation, 61–63 intracrine signaling mechanism inhibitory pathway, 70–72 stimulatory pathway, 66–70 mouse embryo, 48–50 receptors, 3–5 schematics, 48 stem cell pluripotency, 50–51 type I receptor kinase domain structure, 31 Transmembrane prostate androgen-induced RNA, 94 Tumor cell migration, activins breast cancer, 135–137
colon cancer, 138 prostate cancer, 134–135 Two-hit hypothesis, 332, 333 Type 2 diabetes mellitus, 326–327 Type I BMP receptors, 109–110 Type II BMP receptors, 110 Type IV collagenase production, 241–242 U 5’ Untranslated regions (UTRs), inhibin, 151, 152 W White adipose tissue, 223–224
