Yen & Jaffe's Reproductive Endocrinology: Physiology, Pathophysiology, and Clinical Management [9 ed.] 9780323810074

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REPRODUCTIVE ENDOCRINOLOGY

Physiology Pathophysiology, and Clinical Management

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STRAUSS BARBIERI DOKRAS WILLIAMS WILLIAMS

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Yen & Jaffe’s

REPRODUCTIVE ENDOCRINOLOGY

Ninth Edition

Yen & Jaffe’s

REPRODUCTIVE ENDOCRINOLOGY

Physiology, Pathophysiology, and Clinical Management

JEROME F. STRAUSS III, MD, PHD Emeritus Professor Department of Obstetrics and Gynecology Perelman School of Medicine University of Pennsylvania Philadelphia, Pennsylvania United States

ROBERT BARBIERI, MD

Kate Macy Ladd Distinguished Professor Obstetrics, Gynecology and Reproductive Biology Harvard Medical School; Chair, Emeritus Department of Obstetrics and Gynecology Brigham and Women’s Hospital Boston, Massachusetts United States

ANUJA DOKRAS, MD, MHCI, PHD Professor Obstetrics and Gynecology University of Pennsylvania Philadelphia, Pennsylvania United States

CARMEN J. WILLIAMS, MD, PHD Chapel Hill, North Carolina United States

ZEV WILLIAMS, MD, PHD REI Division Chief Obstetrics and Gynecology Columbia University Medical Center New York, New York United States

Elsevier 1600 John F. Kennedy Blvd. Ste 1800 Philadelphia, PA 19103-­2899 YEN & JAFFE’S REPRODUCTIVE ENDOCRINOLOGY: PHYSIOLOGY, PATHOPHYSIOLOGY, AND CLINICAL MANAGEMENT, NINTH EDITION Copyright © 2024 by Elsevier Inc. All rights reserved

ISBN: 978-0-323-81007-4

No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein).

Notice Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds or experiments described herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made. To the fullest extent of the law, no responsibility is assumed by Elsevier, authors, editors or contributors 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.

Previous editions copyrighted 2019, 2014, 2009, 2004, 1999, 1991, 1986, and 1978.

Executive Content Strategist: Nancy Duffy Senior Content Development Specialist: Joanie Milnes Publishing Services Manager: Shereen Jameel Project Managers: Nadhiya Sekar/Manikandan Chandrasekaran Design Direction: Bridget Hoette Printed in India Last digit is the print number:

Contributors Richard J. Auchus, MD, PhD

Professor Division of Metabolism, Endocrinology, and Diabetes Department of Internal Medicine University of Michigan Ann Arbor, Michigan United States Robert Barbieri, MD

Kate Macy Ladd Distinguished Professor Obstetrics, Gynecology and Reproductive Biology Harvard Medical School; Chair, Emeritus Department of Obstetrics and Gynecology Brigham and Women’s Hospital Boston, Massachusetts United States Misty Blanchette Porter, MD, FAIUM

Vice Chair for Education and Faculty Affairs Obstetrics, Gynecology and Reproductive Sciences Larner College of Medicine Burlington, Vermont United States Kassie Bollig, MD

Assistant Professor Department of Obstetrics and Gynecology Penn State College of Medicine and Penn State Health Hershey, Pennsylvania United States Paula C. Brady, MD

Assistant Professor Obstetrics and Gynecology Columbia University Irving Medical Center New York, New York United States Robert Brannigan, MD

Professor and Vice Chair of Clinical Urology Director of Andrology Fellowship Urology Northwestern University, Feinberg School of Medicine Chicago, Illinois United States Leah H. Bressler, MD

Department of Obstetrics and Gynecology University of North Carolina—Chapel Hill Chapel Hill, North Carolina United States

Serdar E. Bulun, MD

Chair Obstetrics and Gynecology Northwestern University Feinberg School of Medicine Chicago, Illinois United States

Anuja Dokras, MD, MHCI, PhD

Professor Obstetrics and Gynecology University of Pennsylvania Philadelphia, Pennsylvania United States Daniel A. Dumesic, MD

Professor of Endocrinology Endocrine Unit University of Palermo School of Medicine Palermo, Italy

Professor Department of Obstetrics and Gynecology University of California—Los Angeles Los Angeles, California United States

Alice Y. Chang, MD

Francesca E. Duncan, PhD

Enrico Carmina, MD

Endocrinology Mayo Clinic Jacksonville, Florida United States R. Jeffrey Chang, MD

Professor Emeritus Obstetrics, Gynecology and Reproductive Sciences University of California—San Diego La Jolla, California United States John A. Cidlowski, PhD

Senior Investigator Laboratory of Signal Transduction National Institute of Environmental Health Sciences, NIH, Research Triangle Park, North Carolina United States Amander T. Clark, PhD

Professor Molecular, Cell and Developmental Biology UCLA Center for Reproductive Science, Health and Education Broad Stem Cell Research Center University of California—Los Angeles Los Angeles, California United States Emmanuèle C. Dêlot, PhD

Research Professor George Washington University Center for Genetic Medicine Research Children’s National Research and Innovation Campus Washington, Washington DC United States James A. Dias, BS, MS, PhD

Vice President for Research Emeritus State University of New York; Professor Emeritus Department of Biomedical Science Albany, New York United States

Assistant Professor Obstetrics and Gynecology Northwestern University Chicago, Illinois United States

Andrea G. Edlow, MD, MSc

Staff Physician, Obstetrics and Gynecology, Division of Maternal-­Fetal Medicine Massachusetts General Hospital; Associate Professor Obstetrics, Gynecology and Reproductive Biology Harvard Medical School Boston, Massachusetts United States Dieter Egli, PhD

Associate Professor Department of Pediatrics and Obstetrics & Gynecology Columbia Stem Cell Initiative Columbia University Irving Medical Center New York, New York United States William S. Evans, MD

Professor Emeritus of Medicine School of Medicine University of Virginia Charlottesville, Virginia United States Bart C.J.M. Fauser, MD, PhD

Professor Emeritus Reproductive Medicine University of Utrecht and University Medical Center Utrecht, The Netherlands Jill P. Ginsberg, MD

Professor Pediatrics Perelman School of Medicine Director, Cancer Survivorship Pediatric Oncology Children’s Hospital of Philadelphia Philadelphia, Pennsylvania United States

v

vi

Contributors

Elizabeth S. Ginsburg, MD

Professor, Harvard Medical School Obstetrics and Gynecology Brigham and Women’s Hospital Boston, Massachusetts United States Linda C. Giudice, MD, PhD

Distinguished Professor Obstetrics, Gynecology and Reproductive Sciences University of California—San Francisco San Francisco, California United States Sierra Goldsmith, MS

Infertility Center of St. Louis St. Lukes Hospital St. Louis, Missouri United States Steven Robert Goldstein, MD

Professor Obstetrics and Gynecology New York University Grossman School of Medicine New York, New York United States Clarisa R. Gracia, MD, MSCE

Professor Director Reproductive Endocrinology and Infertility Department of Obstetrics and Gynecology University of Pennsylvania Philadelphia, Pennsylvania United States Janet E. Hall, MD, MSc

Clinical Director and Senior Investigator National Institute of Environmental Health Sciences, NIH, Research Triangle Park, North Carolina United States Eduardo Hariton, MD, MBA

Reproductive Endocrinology and Infertility Physician Reproductive Science Center of the San Francisco Bay Area VP of Strategic Initiatives and Managing Director of the USF Innovation Fund US Fertility Oakland, California United States Katsuhiko Hayashi, PhD

Professor Stem Cell Biology and Medicine Faculty of Medical Sciences Kyushu University Fukuoka, Japan

Xin He, MD

Professor Division of Metabolism, Endocrinology, and Diabetes Department of Internal Medicine University of Michigan Ann Arbor, Michigan United States Rinath Jeselsohn, MD

Assistant Professor Medical Oncology DFCI Boston, Massachusetts United States Daniel J. Kaser, MD

Physician, Director of Third Party Reproduction and LGBTQ+ Care Reproductive Endocrinology & Infertility Reproductive Medicine Associates of Northern California San Francisco, California United States Andrew M. Kelleher, PhD

Department of Obstetrics Gynecology and Women’s Health University of Missouri Columbia, Missouri United States Laxmi A. Kondapalli, MD, MSCE

Physician Reproductive Endocrinology and Infertility Colorado Center for Reproductive Medicine Lone Tree, Colorado United States

Peter Y. Liu, MBBS, PhD

Professor David Geffen School of Medicine University of California – Los Angeles; Investigator The Lundquist Institute at Harbor-UCLA Medical Center Los Angeles, California United States Roger A. Lobo, MD

Professor Obstetrics and Gynecology Columbia University College of Physicians & Surgeons New York, New York United States Thanh-Ha Luu, MD

Invia Fertility Chicago, Illinois United States

Philip Marsh, MS, TS

Embryologist Obstetrics and Gynecology University of California—San Francisco San Francisco, California United States John C. Marshall, MD, PhD

Andrew D Hart Professor of Medicine Emeritus Division of Endocrinology and Metabolism Department of Medicine University of Virginia Charlottesville, Virginia United States Christopher R. McCartney, MD

Clinical Professor of Obstetrics and Gynecology Vanderbilt University Medical Center Managing Partner Fertility Associates of Memphis Memphis, Tennessee United States

Professor of Medicine Department of Medicine Division of Endocrinology and Metabolism Center for Research in Reproduction University of Virginia School of Medicine Charlottesville, Virginia United States

Monica M. Laronda, PhD

Melissa Menezes, MD

Richard S. Legro, MD

Sam Mesiano, PhD

William Hanna Kutteh, MD, PhD, HCLD

Assistant Professor Pediatrics Lurie Children’s Hospital Northwestern University Chicago, Illinois United States

Chair Department of Obstetrics and Gynecology; Professor Obstetrics and Gynecology and Public Health Sciences Penn State College of Medicine and Penn State Health Hershey, Pennsylvania United States

Assistant Clinical Professor of Pediatrics Division of Adolescent Medicine Department of Pediatrics Children’s Hospital at Montefiore Bronx, New York United States William H Weir MD Professor of Reproductive Biology Department of Reproductive Biology Case Western Reserve University; Vice Chair for Research Department of Obstetrics and Gynecology University Hospitals of Cleveland Cleveland, Ohio United States

Contributors

Diana Monsivais, PhD

Department of Pathology & Immunology Baylor College of Medicine Houston, Texas United States Jerrine Morris, MD, MPH

Third Year Clinical Fellow REI Obstetrics, Gynecology and Reproductive Sciences University of California—San Francisco San Francisco, California United States Prema Narayan, PhD

Associate Professor Physiology Southern Illinois University School of Medicine Carbondale, Illinois United States Ralf Nass, MD

Assistant Professor of Medicine School of Medicine University of Virginia Charlottesville, Virginia United States Kathleen O’Neill, MD, MSTR

Assistant Professor Obstetrics and Gynecology University of Pennsylvania Philadelphia, Pennsylvania United States Sharon E. Oberfield, MD

Professor of Pediatrics and Division Director Pediatric Endocrinology, Diabetes and Metabolism Columbia University Medical Center New York, New York United States Takehiko Ogawa, MD, PhD

Professor Department of Regenerative Medicine Graduate School of Medicine Yokohama City University Yokohama, Japan Giovanna Olivera, MS, TS

Senior Embryologist Obstetrics and Gynecology/REI University of California—San Francisco San Francisco, California United States Kyle E. Orwig, PhD

Professor Obstetrics, Gynecology and Reproductive Sciences Magee-Womens Research Institute University of Pittsburgh School of Medicine Pittsburgh, Pennsylvania United States Stephanie A. Pangas, BA, MS, PHD

Associate Professor Pathology & Immunology Baylor College of Medicine Houston, Texas United States

Alex J. Polotsky, MD

Medical Director SGF Colorado; Professor Obstetrics and Gynecology, Reproductive Endocrinology and Infertility Division and Fellowship; Director University of Colorado School of Medicine Greenwood Village, Colorado United States Molly Quinn, MD

Assistant Professor Department of Obstetrics and Gynecology Division of Reproductive Endocrinology and Infertility Keck School of Medicine of the University of Southern California Los Angeles, California United States Catherine Racowsky, PhD

Professor Emeritus Obstetrics, Gynecology and Reproductive Biology Brigham & Women’s Hospital and Harvard Medical School Boston, Massachusetts United States Lauren Kendall Rauchfuss, MD

Obstetrics and Gynecology Mayo Clinic Rochester, Minnesota United States

Salustiano Ribeiro, MS, TS, ELS/ALS

Embryologist Obstetrics, Gynecology and Reproductive Sciences University of California—San Francisco San Francisco, California United States Jessica Rieder, MD, MS

Associate Clinical Professor of Pediatrics Division of Adolescent Medicine Department of Pediatrics Children’s Hospital at Montefiore Bronx, New York United States Tamar Reisman, MD

Assistant Professor of Medicine Division of Endocrinology and Center for Transgender Medicine and Surgery Icahn School of Medicine at Mount Sinai New York, New York United States Andrea H. Roe, MD, MPH

Assistant Professor Department of Obstetrics and Gynecology Perelman School of Medicine University of Pennsylvania

vii

Cassandra Roeca, MD

Assistant Professor Obstetrics and Gynecology University of Colorado School of Medicine Shady Grove Fertility Denver, Colorado United States Andrew Runge, BS, TS

Senior Embryologist Obstetrics, Gynecology and Reproductive Sciences University of California—San Francisco San Francisco, California United States Mitchell Rosen, MD, HCLD

Professor Obstetrics, Gynecology and Reproductive Sciences University of California—San Francisco San Francisco, California United States Joshua D. Safer, MD, FACP, FACE

Executive Director Center for Transgender Medicine and Surgery Mount Sinai Health System; Professor of Medicine Icahn School of Medicine at Mount Sinai New York, New York United States Nanette Santoro, MD

Professor and Chair Obstetrics and Gynecology University of Colorado School of Medicine Aurora, Colorado United States Karen Schindler, PhD

Associate Professor Department of Genetics Human Genetics Institute of NJ Rutgers The State University of New Jersey New Brunswick, New Jersey United States Peter N. Schlegel, MD

James J. Colt Professor Urology Weill Cornell Medicine New York, New York United States Courtney A. Schreiber, MD, MPH

Stuart and Emily B.H. Mudd Professor of Human Behavior & Reproduction Chief, Division of Family Planning Department of Obstetrics and Gynecology Executive Director, FOCUS on Health and Leadership for Women Perelman School of Medicine University of Pennsylvania Philadelphia, Pennsylvania



viii

Contributors

Danny J. Schust, MD

Professor and Vice Chair for Research Director, Duke Reproductive Endocrinology and Infertility Fellowship Training Program Department of Obstetrics and Gynecology Duke University Durham, North Carolina United States

Aleksandar K. Stanic, MD, PhD

Associate Professor Department of Obstetrics and Gynecology Division of Reproductive Endocrinology and Infertility, Reproductive Sciences University of Wisconsin-Madison Madison, Wisconsin United States

Eric Vilain, MD, PhD

Associate Vice Chancellor Clinical and Translational Science Professor Department of Pediatrics University of California Irvine, California United States Shannon Whirledge, PhD, MHS

Professor and Consultant, Obstetrics and Gynecology Mayo Clinic Rochester, Minnesota United States

Associate Professor Department of Obstetrics, Gynecology and Reproductive Sciences Yale School of Medicine New Haven, Connecticut United States

Jerome F. Strauss III, MD, PhD

Carmen J. Williams, MD, PhD

Clinical and Research Fellow Obstetrics and Gynecology Massachusetts General Hospital Boston, Massachusetts United States

Professor Emeritus Department of Obstetrics and Gynecology Perelman School of Medicine, University of Pennsylvania Philadelphia, Pennsylvania United States

Sherman J. Silber, MD

Yousin Suh, PhD

Zev Williams, MD, PhD

Thalia R. Segal, MD

Assistant Professor Obstetrics, Gynecology, and Reproductive Sciences University of California—San Francisco San Francisco, California United States Molly R. Siegel, MD

Director Infertility Center of St. Louis St. Lukes Hospital St. Louis, Missouri United States Peter J. Snyder, MD

Professor of Medicine Medicine University of Pennsylvania Philadelphia, Pennsylvania United States Aviva B. Sopher, MD, MS

Associate Professor Department of Pediatrics Columbia University Irving Medical Center New York, New York United States Thomas E. Spencer, PhD

Professor Obstetrics Gynecology and Women’s Health University of Missouri Columbia, Missouri United States Frank Z. Stanczyk, PhD

Professor Department of Obstetrics and Gynecology Department of Population and Public Health Sciences Keck School of Medicine University of Southern California Los Angeles, California United States

Elizabeth A. Stewart, MD

Charles and Marie Robertson Professor of Reproductive Sciences Professor of Genetics and Development Director of Reproductive Aging Program Columbia University Vagelos College of Physicians & Surgeons New York, New York United States Alok K. Tewari, MD, PhD

Physician Medical Oncology Dana-­Farber Cancer Institute; Instructor in Medicine Harvard Medical School Boston, Massachusetts United States Nicholas A. Tritos, MD, DSc, MMSc

Staff Neuroendocrinologist Neuroendocrine Unit Massachusetts General Hospital; Associate Professor of Medicine, Medicine Harvard Medical School Boston, Massachusetts United States Jenna Turocy, MD

Assistant Professor Obstetrics and Gynecology Columbia University Irving Medical Center New York, New York United States Alfredo Ulloa-­Aguirre, MD, DSc

Scientific Director RAI, Instituto Nacional de Ciencias Médicas y Nutrición SZ-Universidad Nacional Autónoma de México Mexico DF, Mexico

Senior Investigator Reproductive & Developmental Biology Laboratory National Institute of Environmental Health Sciences Research Triangle Park, North Carolina United States REI Division Chief Obstetrics and Gynecology Columbia University Medical Center New York, New York United States Selma Feldman Witchel, MD

Professor of Pediatrics Division of Pediatric Endocrinology, Department of Pediatrics UPMC Children’s Hospital of Pittsburgh, University of Pittsburgh Pittsburgh, Pennsylvania United States Tracey J. Woodruff, PhD, MPH

Professor and Director Program on Reproductive Health and the Environment Department of Obstetrics, Gynecology, and Reproductive Sciences Institute for Health Policy Studies University of California San Francisco, California United States Steven L. Young, MD, PhD

Professor Reproductive Endocrinology and Infertility, Obstetrics, and Gynecology University of North Carolina School of Medicine Chapel Hill, North Carolina United States Marya G. Zlatnik, MD, MMS

Professor Obstetrics, Gynecology and Reproductive Sciences University of California—San Francisco San Francisco, California United States

Preface The first edition of Yen and Jaffe, Reproductive Endocrinology was published in 1978, the year of the first birth conceived through in-­vitro fertilization (IVF). The foreword to the edition was written by Roger Guillemin MD PhD, who received the 1977 Nobel Prize in Physiology or Medicine for contributions to the discovery of thyrotropin releasing hormone and gonadotropin releasing hormone. In the foreword, Dr Guillemin highlighted the importance of advances in neuroendocrinology to our understanding of reproduction. For the fifth edition of the text, published in 2004, the founding editors transitioned editorial responsibility to us. At the time, the importance of the field of reproductive endocrinology and infertility was firmly established. Scientific advances were rapidly occurring in gamete biology and the genetics of reproduction, and IVF was established as a highly effective and safe treatment for infertility. Acknowledging the importance of IVF and the field of reproductive endocrinology, Robert G Edwards PhD was awarded the 2010 Nobel Prize in Physiology or Medicine for his contributions to the development of IVF. The publication of the ninth edition of Yen & Jaffe marks another milestone in the history of this text, which has achieved a prominent place in the field of reproductive medicine. We were entrusted with the future of the book by its originators, who were giants in our field. Having done our best to meet their expectations, we now prepare for another transition in editorship of the text. With the publication of the ninth edition, we will be transitioning editorial responsibilities to Anuja Dokras, MD, MHCI, PhD, Carmen J. Williams, MD, PhD, and Zev Williams, MD, PhD, our co-­editors of this edition. Collectively, their expertise covers the breadth of science underpinning reproductive

endocrinology and infertility, and we are confident that they will ensure that the text will continue to be comprehensive and authoritative. We are extremely grateful to our many colleagues who authored chapters in the five editions we have overseen. We are especially appreciative of the efforts of those who contributed chapters to the ninth edition in the face of challenges resulting from a pandemic that disrupted clinical care, academic pursuits, and training. The assistance we have received from the professional staff at Elsevier, who facilitated the publication process for the 20 years during which we served as editors, has been invaluable, and we want to thank Nancy Duffy, Joanie Milnes, Nadhiya Sekar, Manikandan Chandrasekaran for their outstanding support in the preparation of the ninth edition. Yen & Jaffe has always been an evolving text, and that will continue to be one of its hallmarks. The ninth edition incorporates new chapters that cover advances in basic and clinical science that have enriched the field since publication of the previous edition, including emerging technologies involving gamete production and maturation, genetic testing and genetic manipulation, and uterus transplantation. Other chapters have been extensively revised to cover contemporary practices in assisted reproduction, fertility preservation, and ovulation induction. We have been honored to carry on the work initiated by Sam Yen and Bob Jaffe and will always be mindful of their substantive scientific contributions and the opportunity they gave to us to carry forward their tradition of excellence. Jerome F. Strauss III, MD, PhD Robert Barbieri, MD

ix

Robert B. Jaffe, MD

1933–2020 In 2020, endocrinology in general—and reproductive endocrinology in particular—lost a giant in basic and translational reproductive endocrinology research. Robert B. Jaffe MD was a visionary leader whose contributions were essential to the development of the field of reproductive endocrinology. Following training at the University of Michigan, the University of Colorado, and the Karolinska Institute with Egon Diczfalusy, Dr Jaffe joined the Michigan faculty, rising to professor over a nine-­year tenure in Ann Arbor. Drs Jaffe and Samuel S.C. Yen began a productive collaboration during this time. Dr Yen would drive from Cleveland, where he was on the Case Western faculty, to Ann Arbor with biological samples in the trunk of his car and have them analyzed for myriad reproductive hormones in Dr Jaffe’s laboratory. Following Dr Yen’s move to San Diego in 1970 and Dr Jaffe’s move to San Francisco in 1973, the two maintained a close relationship. In 1976, they were both Visiting Scholars of the Rockefeller Foundation working at the Villa Serbelloni, Lake Como, Italy. There, they conceived the idea of Yen & Jaffe’s Reproductive Endocrinology, first published in 1978. The book became one of the pillars of the field of reproductive medicine. It is often referred to as “The Bible,” with the ninth edition in preparation. To fully appreciate the impact of the visionary thinking that gave birth to this textbook, one has to consider its temporal context. Reproductive medicine was very much in its infancy in the mid-­1970s. At that time, the development of methods to quantify physiological levels of proteins and steroid hormones,

xii

Dedication

the elucidation of mechanisms of hormone action, and advances in reproductive biology were transforming the science of endocrinology. Innovation in surgical and medical approaches to the diagnosis and treatment of infertility—and the broader availability of hormones to regulate fertility and stimulate the gonads—were establishing the foundations of a new clinical discipline. The first edition of Yen & Jaffe assembled and synthesized the core knowledge upon which this field, Reproductive Endocrinology and Infertility, would rapidly grow, both as a science and as a subspecialty of Obstetrics and Gynecology. Dr Jaffe was an indefatigable leader and champion for academic reproductive endocrinology. As an example, he founded and was the principal investigator for 25 years of the innovative NIH Reproductive Scientist Development Program (RSDP). The RSDP trained dozens of academic leaders in our field and continues to actively support the research career development of early career investigators in reproduction. Dr Jaffe was an exceptional collaborator. He developed a journal club focused on ovarian biology and cancer at MD Anderson Cancer Center, regularly flying from San Francisco to Houston to help lead the discussion of seminal publications. This remarkable collaboration resulted in an NIH-­funded research project involving both UCSF and MD Anderson. Dr Jaffe received many honors and awards, including the Distinguished Scientist Award from the American Society for Reproductive Medicine and the Sydney H. Ingbar Distinguished Service Award from the Endocrine Society. He was a President of the Endocrine Society’s Hormone Foundation and a member of the National Academies of Sciences, Engineering, and Medicine. Dr Jaffe will be greatly missed by his many friends and colleagues throughout the world. His legacy will live on through his many trainees and this book.

Contents Video Contents

xv

PART 1 The Fundamentals of Reproduction 1.

Neuroendocrinology of Reproduction

1 1

Christopher R. McCartney and John C. Marshall

2.

The Gonadotropin Hormones and Their Receptors 23 Prema Narayan, Alfredo Ulloa-Aguirre, and James A. Dias

3.

Prolactin in Human Reproduction

Steroid Hormones and Other Lipid Molecules Involved in Human Reproduction 73 Jerome F. Strauss III, Emanuela Ricciotti, and Garret A. FitzGerald

5.

Steroid Hormone Action

17.

110

18.

Growth Factors and Reproduction

125

Diana Monsivais and Stephanie A. Pangas

7.

Neuroendocrine Control of the Menstrual Cycle 142 Janet E. Hall

8.

The Ovarian Life Cycle

19.

158

Meiosis, Fertilization, and Preimplantation Embryo Development 188

Structure, Function, and Evaluation of the Female Reproductive Tract 217 Andrew M. Kelleher, Leah H. Bressler, Steven L. Young, and Thomas E. Spencer

11.

Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development 254 Sam Mesiano

12.

The Breast

277

20.

21.

461

Physiological and Pathophysiological Alterations of the Neuroendocrine Components of the Reproductive Axis 475

22.

Polycystic Ovary Syndrome and Hyperandrogenic States 516 R. Jeffrey Chang, Anuja Dokras, and Daniel A. Dumesic

23.

Female Infertility

548

24.

Male Infertility

575

Peter N. Schlegel

25.

Endocrine Disturbances Affecting Reproduction 587 Xin He, Alice Y. Chang, and Richard J. Auchus

26.

Benign Uterine Diseases and Dysfunction

601

Lauren Kendall Rauchfuss and Elizabeth A. Stewart

27.

Endometriosis

619

Serdar E. Bulun

28.

Recurrent Pregnancy Loss

650

The Hypothalamo-Pituitary Unit, Testis, and Male Accessory Organs 285

Menopause and Aging

Endocrine Diseases of Pregnancy

665

Molly R. Siegel and Andrea G. Edlow

30.

Male Reproductive Aging

712

Transgender Hormonal Treatment

735

Tamar Reisman and Joshua D. Safer

338

Immunology and Reproduction

Hormone-Responsive Cancers Rinath Jeselsohn and Alok K. Tewari

31.

300

Peter J. Snyder

16.

Environmental Factors and Reproduction

Ralf Nass and William S. Evans

Roger A. Lobo and Yousin Suh

15.

449

Linda C. Giudice, Marya G. Zlatnik, Thalia R. Segal, and Tracey J. Woodruff

29.

Peter Y. Liu

14.

Nutrition and Reproduction

Jenna Turocy and Zev Williams

Robert Barbieri

13.

395

Nanette Santoro, Alex J. Polotsky, Thanh-Ha Luu, Melissa Menezes, Jessica Rieder, Laxmi A. Kondapalli, and Cassandra Roeca

Carmen J. Williams and Karen Schindler

10.

Puberty: Gonadarche and Adrenarche

Paula C. Brady and Robert Barbieri

Jerome F. Strauss III and Carmen J. Williams

9.

365

Aviva B. Sopher, Sharon E. Obereld, and Selma Feldman Witchel

Shannon Whirledge and John A. Cidlowski

6.

Differences of Sex Development Emmanuèle C. Délot and Eric Vilain

56

Nicholas A. Tritos

4.

PART 2 Pathophysiology and Therapy: Pediatric, Adolescent, and Adult 365

345

Aleksandar K. Stanic, William Hanna Kutteh, Kassie Bollig, and Danny J. Schust

PART 3 Reproductive Technologies 32.

742

Evaluation of Hormonal Status

742

Enrico Carmina, Frank Z. Stanczyk, and Rogerio A. Lobo

xiii

xiv 33.

Contents

Pelvic Imaging in Reproductive Endocrinology 772

38.

Misty Blanchette Porter and Steven Robert Goldstein

34.

Medical Approaches to Ovarian Stimulation for Infertility 813 Richard S. Legro and Bart C.J.M. Fauser

35.

Assisted Reproduction: Clinical Practice

Monica M. Laronda, Takehiko Ogawa, Sierra Goldsmith, Sherman J. Silber, Amander T. Clark, Katsuhiko Hayashi, and Kyle E. Orwig

39.

842

Daniel J. Kaser, Elizabeth S. Ginsburg, and Catherine Racowsky

36.

Gamete and Embryo Manipulation

876

Mitchell Rosen, Eduardo Hariton, Philip Marsh, Jerrine Morris, Giovanna Olivera, Andrew Runge, Molly Quinn, Salustiano Ribeiro, and Rhodel Simbulan

37.

Fertility Preservation

907

Francesca E. Duncan, Robert Brannigan, Jill P. Ginsberg, and Clarisa R. Gracia

Emerging Technologies: In Vitro-Derived Germ Cells and Gametogenesis 935

Emerging Technologies: Genetic Interventions in the Human Germ Line: Mitochondrial Replacement and Gene Editing 948 Dieter Egli

40.

Emerging Technologies: Uterus Transplantation 955 Kathleen O’Neill

41.

Contraception

970

Andrea H. Roe and Courtney A. Schreiber

Index

985

Video Contents 17.1 26.1

26.2

Developmental anomalies of the uterus and vagina: classification and treatment Overview of the FIGO classification of uterine leiomyoma and the various operative techniques used for myomectomy, including hysteroscopic myomectomy, laparoscopic myomectomy, robotic-­ assisted myomectomy, and open myomectomy Overview of different endometrial pathologies and the various hysteroscopic techniques employed for their resection, including hysteroscopic myomectomy, hysteroscopic metroplasty, and hysteroscopic lysis of adhesions

27.1 33.1 33.2 33.3 36.1 37.1

Descriptions of lesion types, how to report endometriosis findings, and the various surgical management options for each endometriosis scenario Sonohysterography (SHG) dynamic imaging of performance of an SHG with a polyp or myoma Hysterosalpingo-­contrast sonography video Dynamic ultrasound demonstrating anterior and posterior cul-­de-­sac adhesions relative to deeply invasive endometriosis (sliding sign) The IVF laboratory: management and technologies Mini laparotomy, ovarian tissue preparation on the bench prior to vitrification, and ovarian tissue transplantation

  

xv

PART I

1

1

The Fundamentals of Reproduction

Neuroendocrinology of Reproduction Christopher R. McCartney and John C. Marshall

OUTLINE CENTRAL CONTROL OF REPRODUCTION NEUROENDOCRINOLOGY: THE INTERFACE BETWEEN NEUROBIOLOGY AND ENDOCRINOLOGY Anatomy of the Reproductive Hypothalamic-­Pituitary Axis Gonadotropin-­Releasing Hormone: The Final Common Pathway for the Central Control of Reproduction Neuronal Inputs Into Gonadotropin-­Releasing Hormone Neurons Gonadotropin-­Releasing Hormone Pulse Generator Gonadotropin-­Releasing Hormone Secretion During Development and in Adulthood Physiologic Development of Reproductive Neuroendocrine Function Patterns of Pulsatile Gonadotropin-­Releasing Hormone Secretion in Adults Feedback Regulation of Gonadotropin-­Releasing Hormone and Gonadotropin Secretion Reproductive Neuroendocrine Adaptations in Settings of Reduced Energy Availability, Stress, and Lactation Interface Between Reproductive Neuroendocrine Function and Energy Availability Impact of Stress on Reproductive Neuroendocrine Function Lactation and Reproductive Neuroendocrine Function Miscellaneous Physiologic Influences on Gonadotropin-­ Releasing Hormone Secretion

CENTRAL CONTROL OF REPRODUCTION Successful reproduction is essential to the survival of a species. The reproductive system represents a highly complex functional organization of diverse tissues and signaling pathways that, when properly functioning, ensures a number of key endpoints. The most important of these are the adequate production and development of gametes (ova and sperm), successful delivery of gametes for fertilization, and physiologic preparation for possible pregnancy in women. Neuroendocrine systems are the principal drivers of reproductive function in both men and women. In particular, hypothalamic gonadotropin-­releasing hormone (GnRH) is the primary—if not exclusive—feedforward stimulatory signal to gonadotrope cells of the anterior pituitary, which induces the synthesis and secretion of both luteinizing hormone (LH) and follicle-­stimulating hormone (FSH). Together, these two gonadotropins direct the primary functions of the reproductive axis: gamete development and gonadal sex steroid synthesis. Given its critical importance to a species, the reproductive system must be robust, continuing to function properly in the

face of various physiologic perturbations. In contrast, in settings of marked physiologic stress (e.g., significantly reduced energy availability), mechanisms that temporarily limit fertility—the teleological outcome of which is metabolically expensive in women—are biologically advantageous for the individual and, ultimately, the species. Appropriate function (or quiescence) of the reproductive system is governed by a number of intricate relationships. For example, feedback signals from the gonads (e.g., circulating sex steroid concentrations) communicate the status of gonadal function to the hypothalamic-­pituitary axis; these signals in turn influence GnRH and gonadotropin output, rendering a coordinated and tightly regulated feedback system that maintains gonadal function within narrow limits. The reproductive system also has extensive interactions with other neuroendocrine systems, such as those involved with energy balance and adaptations to stress. The reproductive neuroendocrine system integrates these myriad feedback signals, and the GnRH-­secreting neuronal network represents the final common pathway for the central control of reproduction. Thus the regulation of GnRH secretion represents a major focus of reproductive neuroendocrinology. Much of our understanding of reproductive neuroendocrinology has emerged from the study of rodents, ruminants, and nonhuman primates, largely reflecting the ethical boundaries inherent to human research. Because many neurobiological principles are similar among all mammals, these animal studies continue to be indispensable. Nonetheless, certain aspects of reproductive neuroendocrinology may differ markedly among species. Thus, when available, human data will be prioritized throughout this chapter, but animal studies will also be discussed when appropriate—emphasizing nonhuman primate studies when available— recognizing that specific findings may or may not be generalizable to humans. The reader is referred to Chapters 2, 7, 13, 18, and 21 for additional discussion of neuroendocrine physiology and pathophysiology related to reproduction.

NEUROENDOCRINOLOGY: THE INTERFACE BETWEEN NEUROBIOLOGY AND ENDOCRINOLOGY Endocrinology is the study of cell-­to-­cell signaling that occurs via specific chemicals (hormones) traveling through the bloodstream to influence remote targets. The term “neuroendocrinology” refers to the involvement of the central nervous system (CNS) in this process, particularly the hypothalamus. This field of study has traditionally focused on hypothalamic neuron-­derived factors that influence various target tissues either directly, as with the hormones of the neurohypophysis, or indirectly, as with hypothalamic releasing factors that control anterior pituitary hormone secretion. Neuroendocrine systems direct a wide variety of critical biologic processes, such as growth and development, energy and fluid homeostasis, responses to stress, and reproduction. Neurons are highly specialized, morphologically diverse cells that transmit information via electrical impulses called action potentials. Neurons have a cell body (perikaryon) containing the

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PART I  The Fundamentals of Reproduction

Dendrites Cell body (perikaryon)

Axon Axon terminals Fig. 1.1 Morphologic components of a neuron.

cell nucleus, mitochondria, and synthetic organelles. Neurons also have cell processes that participate in the reception and delivery of electrical impulses (Fig. 1.1). Dendrites are short processes—often extensively branched to increase surface area—that typically receive information by way of afferent electrical impulses from other neurons. The axon is a single neuronal extension that generally transmits efferent electrical impulses away from the cell body via a process called neuronal firing. However, GnRH neuron fibers extending from the cell body to the median eminence (the location of GnRH release) in mice demonstrate characteristics of both axons and dendrites and thus have been called dendrons.1 In unstimulated neurons, the inner portion of the neuronal membrane is negatively charged compared with the outer membrane surface; this resting membrane potential is typically between −50 and −75 mV in GnRH neurons. Such electrical polarization reflects transmembrane ionic differences, which are maintained by protein channels that govern transmembrane passage of specific ions (e.g., sodium, potassium, chloride). Regulated changes of transmembrane ion differences may cause the membrane potential to become more or less negative (hyperpolarization and depolarization, respectively). Depolarization to a certain threshold results in a rapid and temporary reversal of membrane potential—an action potential—which is propagated along the neuronal membrane. Notably, the amplitude of the action potential does not vary with the strength of stimulation; instead, once a threshold is reached, a full action potential occurs (the so-­called all-­or-­none phenomenon). However, the degree of neuronal stimulation can alter the frequency of action potentials generated. In this way, neurons transmit information to other neurons and effector tissue cells. Neuronal signals are transferred across neuron-­ to-­ neuron connections (synapses) via chemical neurotransmitters. This process begins with bursts of neuronal firing, which result in the opening of voltage-­gated calcium channels at the axon terminal. The influx of calcium promotes exocytosis of neurotransmitter-­ containing synaptic vesicles, releasing neurotransmitters into the synaptic cleft. Neurotransmitters then activate specific ligand-­ dependent ion channels in the postsynaptic membrane, which can stimulate an action potential in the postsynaptic neuron. A wide variety of factors serve as neurotransmitters, including amino acids (e.g., acetylcholine, glutamate, γ-­aminobutyric acid [GABA]), biogenic amines (e.g., norepinephrine, epinephrine, dopamine, serotonin), and neuropeptides (e.g., kisspeptin, neurokinin B [NKB], dynorphin, β-­endorphin, somatostatin, proopiomelanocortin [POMC], neuropeptide Y [NPY]). Bursts of neuronal firing can also elicit release of neuronal products into the bloodstream to influence remote targets (i.e., neurosecretion of neurohormones). Hypophysiotropic neurons are specialized hypothalamic neurons that secrete peptide-­ releasing factors (GnRH, corticotropin-­ releasing hormone [CRH], thyrotropin-­releasing hormone [TRH], and growth hormone–releasing hormone [GHRH]) into the hypophyseal por tal circulation. These releasing factors in turn stimulate specific

anterior pituitary cell populations. In contrast, hypothalamic release of dopamine into the portal circulation provides tonic inhibition of pituitary prolactin secretion. Hypothalamic neurosecretion of vasopressin and oxytocin, which are released directly into the systemic circulation, alter the function of distant targets such as the renal tubules and uterus, respectively. Neuroglial cells (e.g., astrocytes, ependymal cells, oligodendrocytes, microglia) represent approximately 90% of cells in the CNS. Neuroglia do not conduct action potentials, but they perform critical supportive functions. For example, astrocytes form the supportive framework of the CNS, help isolate synaptic junctions to prevent nonspecific spread of neuronal impulses, facilitate nutrient delivery to neurons, and contribute to the blood-­brain barrier. Of interest, astrocytes have been implicated in the control of GnRH secretion and the mechanisms underlying pubertal onset.2 For example, astrocytes may impact neuronal activity via secretion of numerous growth factors, and astrocytes abundantly appose GnRH neurons; these contacts can influence synaptic input and may be influenced by estrogen in both rodents and nonhuman primates. Similarly, specialized ependymal cells (tanycytes) in the median eminence appear to modify access of GnRH neuron terminals to the hypophyseal portal system.

Anatomy of the Reproductive Hypothalamic-­Pituitary Axis • GnRH neuronal cell bodies are located in the infundibular (arcuate) nucleus and the medial preoptic area of the hypothalamus. • GnRH neurons extend processes to the median eminence, where GnRH gains access to the hypophyseal portal system. • The hypophyseal portal circulation represents the functional connection between hypothalamic GnRH neurons and the gonadotropes of the anterior pituitary.    Portions of the hypothalamus and the anterior pituitary gland constitute the primary effector arm of the central reproductive axis. In particular, hypothalamic neural systems regulate GnRH release into the hypophyseal portal veins, with GnRH being the signal to gonadotropes (anterior pituitary) to secrete LH and FSH. In turn, these gonadotropins direct gonadal (ovarian and testicular) function.

Hypothalamus The hypothalamus is located at the base of the brain (Fig. 1.2). Although small (approximately 10 g, less than 1% of total brain weight), it performs critical functions for maintenance of whole-­ organism homeostasis, including regulation of hunger and body weight, growth, various aspects of metabolism, thirst and renal water handling, body temperature, autonomic function, sleep, circadian rhythms, and emotion. Importantly, the hypothalamus is also a primary control center for reproduction and influences sexual behavior. As an anatomic structure, the hypothalamus does not have discrete borders, but it generally forms the floor and inferior-­lateral walls of the third ventricle (Fig. 1.3). The medial portions of the hypothalamus are primarily made up of cell bodies, whereas the lateral portions are mostly composed of neuron fibers (axons), such as those connecting the medial hypothalamus to other areas of the brain. By convention, closely associated collections of neuron cell bodies are called nuclei; and the paraventricular, dorsomedial, ventromedial, and infundibular nuclei contain a majority of the neurons that secrete hypophysiotropic hormones into the portal circulation. (The human infundibular nucleus is the analogue to the arcuate nucleus in lower mammalian species.) GnRH cell bodies do not form discrete nuclei but are instead diffusely located throughout the preoptic area and the mediobasal

CHAPTER 1  Neuroendocrinology of Reproduction Corpus callosum

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Fig. 1.2 Cross-­sectional representation of the human brain (sagittal plane), including hypothalamus, median eminence, and pituitary gland. (Modified from Johnson MH, Everitt BJ. Essential Reproduction, ed 5, Blackwell Science; 2000:Fig. 6.1.)

Lateral hypothalamus Mammillary nuclear complex

Ventromedial nucleus Infundibular (arcuate) nucleus

Fig. 1.3 Nuclei and areas of the hypothalamus.  (A) By custom, the nuclei and areas of the hypothalamus are often divided into three groups according to their location along the anteroposterior plane: the anterior group, the tuberal group, and the posterior (or mammillary) group. The anterior group is formed by the paraventricular, supraoptic, and suprachiasmatic nuclei, along with the anterior hypothalamic and preoptic areas. The tuberal group—so-­called because of its position above the tuber cinereum, from which the infundibulum or pituitary stalk extends, contains the dorsomedial, ventromedial, and infundibular (arcuate) nuclei along with the median eminence. Along with the paraventricular nucleus, the nuclei of the tuberal group contain a majority of the neurons that secrete hypophysiotropic hormones (i.e., hypothalamic hormones regulate hormone synthesis and release from cells in the anterior pituitary). Finally, the posterior group includes the posterior hypothalamic nucleus and mammillary nuclei. (B) Cross-­sectional representations (coronal planes) of the rostral (1), mid (2), and caudal (3) portions of the human hypothalamus. ([B] Modified from Johnson MH, Everitt BJ. Essential Reproduction, ed 5, Blackwell Science; 2000:Fig. 6.3.)

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PART I  The Fundamentals of Reproduction Tanycytes

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Vein Fig. 1.4 Anatomic relationship between hypothalamic gonadotropin-­releasing hormone (GnRH) neurons and their target cell populations in the adenohypophysis (anterior pituitary). The majority of GnRH neuron cell bodies are located in the infundibular (arcuate) nucleus and the medial preoptic area. GnRH neuron projections terminate at the median eminence, where GnRH is secreted into the hypophyseal portal system. (Modified from Johnson MH, Everitt BJ. Essential Reproduction, ed 5. Blackwell Science; 2000:Fig. 6.4.)

hypothalamus (Fig. 1.4); the latter is situated caudal to the preoptic area, extending from the retrochiasmatic area (i.e., the area located behind the optic chiasm) to the mammillary bodies, and including both the infundibular (arcuate) nucleus and the median eminence.

Median Eminence Positioned at the base of the third ventricle, the median eminence is part of the anatomic link between the hypothalamus and anterior pituitary. The internal zone of the median eminence is located along the ventral floor of the third ventricle and is largely composed of axonal fibers from both magnocellular neurons (larger neurons that secrete vasopressin and oxytocin) and hypophysiotropic neurons as they travel from hypothalamic nuclei/areas to their final destinations; the neurohypophysis (posterior pituitary) and the external zone of the median eminence, respectively (Fig. 1.5). The external zone contains hypophysiotropic neuron terminals, which release hypophysiotropic hormones into an extensive capillary plexus (the proximal end of the hypophyseal portal system). Some nerve terminals in this zone act on other nerve terminals to influence hormone release (e.g., kisspeptin neurosecretion at GnRH neuron terminals influences GnRH release). The ependymal layer lining the third ventricle includes a population of specialized ependymal cells called tanycytes, which have a short process extending toward the ventricular surface and a long process extending into the median eminence toward areas around portal capillaries. The latter tanycyte projections envelop or retract from GnRH nerve terminals during episodes of low and high GnRH neuronal activity, respectively. Thus, tanycytes may influence GnRH secretion via the regulated process of physically isolating GnRH neuron terminals from portal capillaries.3 Tanycytes may also represent a link between cerebrospinal fluid and the external zone of the median eminence (e.g., by transport ing substances from the third ventricle to portal blood).

Portal capillary plexus

Fig. 1.5 Diagram of the median eminence.

The median eminence is among the so-­called circumventricular organs, which lie adjacent to the ventricular system and represent openings in the blood-­brain barrier. Although lipid-­soluble molecules can diffuse in and out of the CNS relatively easily, and cellular transport mechanisms allow selective entry of ions, the blood-­brain barrier functions to protect certain regions of the brain and hypothalamus from larger charged molecules, with physical protection provided by (1) tight junctions between endothelial cells and (2) neuron-­capillary separation by both astrocyte foot processes and microglia. However, the CNS requires feedback signals, including hormonal, metabolic, and toxic cues via macromolecules of peripheral origin that would otherwise be excluded by the blood-­brain barrier. Accordingly, capillaries of the circumventricular organs are fenestrated and permit transcapillary exchange of larger charged molecules (e.g., proteins, peptide hormones). Thus the median eminence represents a key access point for central sensing of peripheral cues. Similarly, fenestrated vessels readily allow entry of hypothalamic-­releasing factors into portal blood.

Hypophyseal Portal Circulation No direct neuronal connections exist between the hypothalamus and the anterior pituitary. However, the hypophyseal portal circulation (hypothalamic-­hypophyseal portal system, pituitary portal system) represents the functional connection between the median eminence and the anterior pituitary (see Fig. 1.4). The superior hypophyseal artery—a branch of the internal carotid artery—subdivides to form an extensive capillary network in the external zone of the median eminence, with loops that reach into the inner zone. Capillary blood then drains into sinusoids that converge into the hypophyseal portal veins. Traversing the pituitary stalk, the hypophyseal portal system forms the primary blood supply of the anterior pituitary. The direction of blood flow is primarily, but not exclusively, from the hypothalamus to the anterior pituitary; some retrograde flow allows for short-­loop hypothalamic feedback.

Pituitary Gland (Hypophysis) The pituitary gland appears as an extension at the base of the hypothalamus and resides cradled within the sella turcica, a saddle-­like structure of the sphenoid bone (see Fig. 1.2). The adenohypophysis (anterior pituitary) is of ectodermal origin, derived from an upward invagination of pharyngeal epithelium (Rathke pouch) during embryologic development. The adenohypophysis is composed of primarily the anterior lobe (pars distalis), which contains specialized cell populations that produce specific hormones: gonadotropes (the gonadotropins LH and FSH), mammotropes (prolactin), corticotropes (adrenocorticotropic

CHAPTER 1  Neuroendocrinology of Reproduction

hormone [ACTH]), thyrotropes (thyroid-­stimulating hormone [TSH]), and somatotropes (growth hormone). The intermediate lobe is vestigial in adult humans but includes a small population of cells (e.g., POMC cells) in contact with the posterior lobe; the pars tuberalis is a slender layer of tissue (e.g., LH-­producing cells and TSH-­producing cells) surrounding the infundibulum (the funnel-­shaped connection between the hypothalamus and the posterior pituitary) and pituitary stalk. In contrast to the adenohypophysis, the neurohypophysis (posterior pituitary) is composed of neural tissue and forms as a downward extension of neuroectodermal tissue from the infundibulum during embryologic development. It is thus a direct extension of the hypothalamus. The neurohypophysis includes the infundibular stalk and the pars nervosa (posterior lobe of the pituitary). The supraoptic and paraventricular nuclei include magnocellular neurons that produce oxytocin and arginine vasopressin (AVP; also known as antidiuretic hormone [ADH]), respectively; these axons project to the posterior lobe of the pituitary, where oxytocin and AVP are secreted into a capillary network that drains into the hypophyseal veins (i.e., directly into the systemic circulation). The posterior lobe also includes specialized glial cells called pituicytes, which envelop or retract from magnocellular nerve terminals during episodes of low and high neuronal activity, respectively.

central reproductive function. However, there are no known parallel or backup pathways for the stimulation of gonadotropin secretion. Thus natural fertility is absolutely dependent on appropriate GnRH secretion. For example, mice with loss-­of-­function variants of the GnRH-­1 gene are hypogonadal, but reproduction can be restored via GnRH-­1 gene therapy6 or transplantation of fetal GnRH neurons.7 Similarly, a variety of human conditions associated with absent (or near-­absent) GnRH secretion lead to pubertal failure, hypogonadotropic hypogonadism, and infertility, all of which can be fully reversed with exogenous GnRH therapy.8 GnRH secretion is influenced by numerous factors, including sex steroids, energy availability, and stress. In some mammalian species, GnRH secretion is also affected by circadian rhythms, photoperiod (e.g., seasonal breeders such as sheep), social cues, and pheromones.

Gonadotropin-­Releasing Hormone Structure GnRH (GnRH-­1 in particular) is a decapeptide, with the amino acid structure (pyro)Glu-­His-­Trp-­Ser-­Tyr-­Gly-­Leu-­Arg-­ Pro-­Gly-­NH2. The amino acid structure of GnRH is identical in essentially all mammalian species; with the exception of the central Tyr-­Gly-­Leu-­Arg segment, the amino acids of GnRH are highly conserved among vertebrate species.9 The GnRH-­1 gene (GNRH1) is located on human chromosome 8 (8p11.2-­p21) and produces a 92–amino acid precursor peptide called prepro-­ GnRH, which includes a signal sequence (23 amino acids), GnRH (10 amino acids), a proteolytic processing site (3 amino acids), and GnRH-­associated peptide (56 amino acids) (Fig. 1.6). The latter peptide can stimulate gonadotropin secretion and inhibit prolactin secretion, although its precise physiologic role, if any, remains unclear. The actions of GnRH are mediated through the GnRH type I receptor. Another form of GnRH (GnRH-­2) and its receptor have been identified in a variety of animal species, including humans.10 GnRH-­2 is a decapeptide with a similar structure to GnRH-­1: (pyro)Glu-­His-­Trp-­Ser-­His-­Gly-­Trp-­Tyr-­Pro-­ Gly-­NH2 (italicized amino acids denote differences compared with GnRH-­1). However, the gene for GnRH-­2 is located on human chromosome 20 (20p13). GnRH-­2 is widely expressed in the CNS and extra-­CNS tissues, and it may contribute to reproductive behavior regulation in some species. In lower animals, GnRH-­2 can act via its own receptor, which is structurally and functionally distinct from the GnRH type I receptor. Although a homologue of the GnRH-­2 receptor gene has

Gonadotropin-­Releasing Hormone: The Final Common Pathway for the Central Control of Reproduction

• Pulsatile GnRH secretion is the proximate stimulus for LH and FSH synthesis and secretion by pituitary gonadotropes. • Although numerous internal and external factors influence gonadotropin secretion via numerous neuronal pathways, GnRH is the final common pathway for the stimulation of LH and FSH release.    GnRH, previously called luteinizing hormone–releasing hormone (LHRH), is synthesized and released by a relatively small population of specialized hypothalamic neurons. GnRH was initially isolated from porcine hypothalami and shown to stimulate pituitary gonadotropin release.4 Although the primary function of GnRH is to regulate pituitary gonadotropin secretion, GnRH also appears to have autocrine and paracrine functions in diverse tissues (e.g., ovary, placenta).5 The regulation of GnRH secretion is complex and involves overlapping pathways, which likely increases the robustness of

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Fig. 1.6 Schematic of gonadotropin-­releasing hormone (GnRH) synthesis.  (A) Representation of prepro-­GnRH, including a 23–amino acid signal sequence, GnRH, a proteolytic processing site (Gly-­Lys-­Arg), and GnRH-­associated peptide. The arrow indicates the site of proteolytic cleavage and C-­amidation. (B) Schematic of neuronal GnRH synthesis and secretion.

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PART I  The Fundamentals of Reproduction

been detected in humans, it includes a frameshift and premature stop codon. Thus the physiologic role of GnRH-­2 in humans remains unclear.

Anatomy of Gonadotropin-­Releasing Hormone-­Secreting Neurons GnRH neurons are a heterogeneous population of hypothalamic neurons. They are relatively few, numbering approximately 7000 in humans, and the majority of GnRH neuronal cell bodies are located in the infundibular (arcuate) nucleus—part of the mediobasal hypothalamus—and the medial preoptic area.11,12 GnRH neurons in the infundibular (arcuate) nucleus appear to be requisite for gonadotropin secretion. For example, selective radiofrequency ablation of the arcuate nucleus in adult female monkeys obliterates gonadotropin secretion.13 Although GnRH neurons are rather loosely affiliated anatomically, they are functionally integrated and form a complex, interconnected network with extensive connections to other neuronal populations. GnRH neurons extend projections through the tuberoinfundibular tract to the median eminence, where neuron terminals gain access to the hypophyseal portal system. Recent work in mice suggests that the GnRH neuron fibers extending from the cell body to the median eminence are morphologically atypical: although they do not exhibit many of the molecular markers classically associated with axons or dendrites, they demonstrate morphologic and functional characteristics of both axons and dendrites, including functional synaptic inputs along the fiber.1,14 Accordingly, the term dendron has been used for such projections.1 In mice, the distal portion of such dendrons exhibit a particularly high density of dendritic spines and synaptic inputs, beyond which the dendron branches into multiple short axons at the median eminence.15 The physiologic function of other GnRH neurons, which arise from the anterior and posterior hypothalamus and project to the limbic system and posterior pituitary, respectively, remains unclear, although some of these circuits may possibly be involved with various behavioral responses.

Fig. 1.7 Gonadotropin-­releasing hormone (GnRH) neuron migration during embryogenesis.  (A) Location of GnRH-­immunoreactive cells (red circles) as a function of embryologic age (mouse). On embryologic day 11 (11E), GnRH cells are located in the nasal (olfactory) placode and presumptive vomeronasal organ (vno). GnRH cells migrate across the cribriform plate toward the olfactory bulb (ob). GnRH neurons then follow the caudal branch of the vomeronasal nerve toward the forebrain and hypothalamus. By day 16 (16E), GnRH neurons largely reside in the preoptic area (poa) of the hypothalamus. (B) Sagittal brain slice (mouse, embryonic day 15) demonstrating the migratory route of GnRH-­immunoreactive cells. Staining is for GnRH and peripherin (a neuronal intermediate filament). BF, Basal forebrain; CP, cribriform plate; gt, ganglion terminale; OB, olfactory bulb; OP/VNO, olfactory placode-­vomeronasal organ. ([A] Modified from Schwanzel-­Fukuda M, Pfaff DW. Origin of luteinizing hormone-­releasing hormone neurons. Nature. 1989;338:161–164; and [B] Modified from Wierman ME, Pawlowski JE, Allen MP, et al. Molecular mechanisms of gonadotropin-­releasing hormone neuronal migration. Trends Endocrinol Metab 2004;15:96–102.)

Embryologic Development of the Gonadotropin-­Releasing Hormone Neuronal Network The ontogeny of GnRH neurons in vertebrate species is unique among neuronal systems of the CNS: nascent GnRH neurons are initially identified outside of the CNS in the nasal placode (sometimes called the olfactory placode). However, GnRH cells migrate during embryologic development, as directly observed in embryonic nasal explant cultures16 and in murine embryonic head slices.17 The specific migratory pathway of GnRH neurons was first demonstrated in mice by documenting the presence of GnRH-­immunoreactive cells in different areas at different stages of embryonic development (Fig. 1.7).18–20 Specifically, GnRH expression is first observed within the nasal placode circa embryonic day 10 or 11. By embryonic day 13, GnRH-­expressing cells are primarily located around the cribriform plate, and GnRH-­ expressing cells begin to reach the hypothalamus by embryonic day 14, approaching their final positions around embryonic day 16. This migratory pathway has been confirmed in both nonhuman primates21 and humans.22 Successful migration of GnRH neurons is inextricably intertwined with olfactory system development, perhaps reflecting the close functional relationship between reproduction and the olfactory system (e.g., pheromones) in mammalian phylogeny. The nasal placode gives rise to nasal epithelium and olfactory sensory neurons, the latter of which extends axonal projections to the olfactory bulb. Vomeronasal neurons are a subset of olfactory neurons believed to be involved with pheromone detection; these axons originate in the vomeronasal organ and largely extend to the accessory olfactory bulb. At the level of the cribriform plate, some olfactory (vomeronasal) axons separate and form a branch that extends caudally into the forebrain. Of great importance, migrating GnRH neurons maintain adhesion to these axons; thus, these olfactory neurons form a critical guidance track for GnRH neuronal migration across the nasal epithelium and through the forebrain toward the hypothalamus.23,24 After reaching the hypothalamus, GnRH neurons detach from olfactory nerve axons and may disperse further before resting. A critical step is the extension

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CHAPTER 1  Neuroendocrinology of Reproduction

Gonadotropin-­Releasing Hormone Neuronal Firing and Gonadotropin-­Releasing Hormone Secretion The activity of many GnRH neurons is marked by bursts of action potentials (burst firing), the patterns and rates of which change across time. Such changes in GnRH neuron firing rates may relate to changes in pulsatile GnRH release at the median eminence, although this hypothesis has not been experimentally verified. Variable firing rate patterns (e.g., times of high and low

firing rates) appear to be intrinsic to GnRH neurons, but they can also be altered by neurotransmitters and neuromodulators (e.g., glutamate, GABA, kisspeptin). Although sex steroids can markedly influence GnRH neuronal firing rates, GnRH neurons lack the primary receptors mediating sex steroid feedback (i.e., estrogen receptor alpha, progesterone receptor, androgen receptor). However, many studies suggest that sex steroid actions on GnRH neuronal activity are mediated primarily via afferent neurons (e.g., those secreting glutamate, GABA, kisspeptin). GnRH neuron cell bodies are relatively scattered across the mediobasal hypothalamus and preoptic area, yet GnRH is secreted into the hypophyseal portal system in a coordinated, pulsatile fashion. Specifically, GnRH secretion is marked by episodic bursts of hormone release into the portal system, as demonstrated in rats,39 sheep,40 and monkeys.41 After being released into the portal vascular compartment, GnRH is rapidly degraded via enzymatic proteolysis, and the half-­life of GnRH in the blood is very short—approximately 2 to 4 minutes. Thus GnRH presentation to gonadotrope cells is intermittent. Pulsatile GnRH secretion is an absolute requirement for long-­ term stimulation of gonadotropin synthesis and secretion, and there is a relatively narrow window of GnRH pulse frequency and amplitude that will optimally stimulate gonadotropin secretion. Intermittent GnRH stimulation of gonadotrope cells can increase (or maintain) GnRH receptors on gonadotropes (the self-­priming or autopriming effect). Thus intermittent GnRH stimulation facilitates or maintains gonadotrope responsiveness to GnRH. However, more frequent exposure to GnRH pulses can reduce gonadotropin responses to GnRH.42 In classic experiments involving rhesus monkeys with hypothalamic lesions that abolished GnRH secretion, once-­hourly exogenous GnRH administration restored pituitary gonadotropin secretion. However, changing from once-­ hourly pulses to twice-­ hourly pulses reduced LH and FSH secretion by 50% to 60%, while 3 to 5 pulses per hour profoundly suppressed plasma LH and FSH concentrations.42 Marked desensitization of gonadotropin release is also observed when changing from once hourly (intermittent) to continuous GnRH administration (Fig. 1.8).43 Although reduced GnRH receptor expression on gonadotropes (i.e., receptor downregulation) plays a role in desensitization, additional mechanisms contribute to the uncoupling of GnRH receptor agonism and gonadotropin synthesis.44

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of GnRH neuronal projections to the median eminence, where GnRH gains access to the hypophyseal portal system. The dependence of GnRH neuronal migration on normal olfactory system development is exemplified by Kallmann syndrome, a form of isolated hypogonadotropic hypogonadism accompanied by an absent or reduced sense of smell (anosmia and hyposmia, respectively).25 In this syndrome, faulty development of the olfactory system renders an inadequate guidance infrastructure for migrating GnRH neurons, leading to absent or incomplete GnRH neuron migration to the hypothalamus. The first identified cause of Kallmann syndrome was a deletion of ANOS1 (formerly the Kallmann syndrome 1 sequence [KAL1] gene), which is located on the X chromosome (Xp22.3) and encodes anosmin-­1, a secreted matrix glycoprotein expressed in the presumptive olfactory bulb. Although precise mechanisms are unclear, anosmin-­1 is believed to be important for the formation of olfactory elements that provide migratory guidance to GnRH neurons as they move out of the nasal placode. Evaluation of a 19-­ week-­ old human fetus with X-­ linked Kallmann syndrome demonstrated GnRH-­immunoreactive cells within a tangle of olfactory and vomeronasal nerves at the dorsal surface of the cribriform plate, along with the absence of olfactory tracts and bulbs.26 In a second human fetus (16 weeks) with X-­linked Kallmann syndrome, GnRH was detected along terminal nerve fascicles in the nasal mucosa only.27 This form of Kallmann syndrome illustrates that without the guidance framework provided by the olfactory neuronal system, GnRH neurons do not appropriately migrate into the hypothalamus and thus cannot release GnRH into the hypophyseal portal system. A number of additional single-­gene defects have been associated with Kallmann syndrome,25 including variants in the genes for prokineticin 2 (PROK2) and its receptor (PROKR2),28 fibroblast growth factor 8 (FGF8) and its receptor fibroblast growth factor receptor 1 (FGFR1),29,30 NMDA receptor synaptonuclear signaling and neuronal migration factor (NSMF; formerly nasal embryonic LH-­releasing hormone factor [NELF]),31 chromodomain helicase DNA binding protein 7 (CHD7),32 semaphorin 3A (SEMA3A),33 and SRY-­box 10 (SOX10).34 The importance of these genes in GnRH neuronal development is corroborated by mouse studies. For example, in fetal mice lacking either Prok2 or Prokr2, GnRH neurons are trapped in a tangled web of olfactory/ vomeronasal axons, with few, if any, reaching the forebrain.35 Although such gene products are clearly important for GnRH neuron migration, their precise roles remain unclear. Of interest, patients with specific Kallmann syndrome-­ related gene variants can exhibit variable penetrance and different phenotypic expressions (even within families), suggesting the importance of other factors such as gene-­gene interactions (oligogenicity).36 Some Kallmann syndrome-­related gene variants (e.g., PROK2, PROKR2, FGF8, FGFR1, NSMF, CHD7) are also associated with normosmic hypogonadotropic hypogonadism.25 In addition, a minority of patients with some Kallmann syndrome-­ related gene variants (e.g., ANOS1, PROKR2, FGFR1, NSMF, or CHD7) may demonstrate partial or full recovery of reproductive function (reversal of hypogonadotropic hypogonadism) in later life,37,38 suggesting plasticity of the GnRH neuronal network and, perhaps, gene-­environment interactions. Mechanisms accounting for such phenomena remain unclear.

0

Days Fig. 1.8 The influence of pulsatile versus continuous gonadotropin-­releasing hormone (GnRH) administration to GnRH-­ deficient monkeys.  Intermittent exogenous GnRH administration reconstitutes normal gonadotropin secretion. However, continuous GnRH infusion leads to a marked reduction (downregulation) of luteinizing hormone (LH; green) and follicle-­stimulating hormone (FSH; purple) concentrations. Resumption of pulsatile GnRH administration restores LH and FSH secretion. (Modified from Belchetz PE, Plant TM, Nakai Y, et al. Hypophysial responses to continuous and intermittent delivery of hypothalamic gonadotropin-releasing hormone. Science.

1

8

PART I  The Fundamentals of Reproduction

GnRH

pGlu His

Trp

Ser

Tyr

Gly

Leu

Arg

Pro

Gly

NH2

Agonists Leuprolide D-amino acid substitution enhances activity Gly Tyr Ser

A

Receptor binding and activation D-amino acid substitution in antagonists

Trp

Leu Arg Pro

His

Gly NH2

NEt

Goserelin

NH2

Histrelin

NH2

Nafarelin

NH2

Triptorelin

NH2

Buserelin Receptor binding only

pGlu

NEt

Antagonists

B

Cetorelix

NH

Ganirelix

NH

Fig. 1.9 Structure of gonadotropin-­releasing hormone (GnRH) and GnRH receptor agonists and antagonists.  (A) Schematic of GnRH-­1 in its folded conformation. Folding around the glycine in position 6 enhances GnRH receptor binding. Substitution of the glycine in position 6 with D-­amino acids stabilizes the molecule in the folded conformation, which increases affinity for the GnRH receptor and reduces metabolic clearance. The amino-­terminal (red) is involved with receptor binding and activation, and GnRH antagonists involve modifications of these residues that prevent receptor activation. The carboxyl-­terminal (green) participates in receptor binding, but not in activation. Substitution at position 10 (e.g., replacement of glycinamide by ethylamide) can increase binding affinity. (B) Amino acid structure of GnRH along with selected GnRH receptor agonists and antagonists. Solid blue circles represent amino acids that are unchanged compared with native GnRH. (From Millar RP, et al. Gonadotropin-­releasing hormone receptors. Endocr Rev. 2004;25:235–275.)

The foregoing phenomenon can be exploited therapeutically with the use of long-­acting GnRH receptor agonists. Such agonists are peptides with structures very similar to that of GnRH but with amino acid substitutions that enhance receptor binding affinity, increase resistance to proteolytic degradation, or both (Fig. 1.9), thus providing continuous GnRH receptor stimulation. Although initial GnRH receptor agonism temporarily (for 1 to 2 weeks) increases gonadotropin release (gonadotropin “flare”), continued agonism leads to desensitization of gonadotropin secretion with accompanying reductions of gonadal sex steroid concentrations to castrate levels (“medical oophorectomy,” “medical castration,” “pseudomenopause”), typically over several weeks. These agents can be useful in the therapy of gonadotropin-­ dependent disorders such as central precocious puberty, endometriosis, and prostate cancer. Peptide GnRH receptor antagonists are also available for clinical use. These antagonists reversibly bind to, but do not stimulate, the GnRH receptor (i.e., competitive antagonism). Thus these agents do not initially stimulate gonadotropin release, and they reduce gonadotropins more rapidly than GnRH agonists— usually within 24 to 72 hours.

Gonadotropin-­Releasing Hormone Stimulation of Gonadotrope Cells The specialized cells that synthesize and secrete gonadotropins (i.e., gonadotropes) are located mainly in the lateral portions of the anterior pituitary gland and constitute approximately 10% of the adenohypophysis cell population. GnRH action at the pituitary gonadotrope begins with GnRH binding to the GnRH type I receptor on the plasma membrane.9 The GnRH type I receptor is a member of the seven-­transmembrane receptor family, a G protein–coupled receptor, and encoded on chromosome 4. GnRH receptor density varies in different physiologic conditions and exhibits a positive correlation with gonadotrope responsiveness to GnRH (e.g., both are high in rodents during preovulatory gonadotropin surges45). GnRH receptor density appears to be modulated primarily by GnRH, with intermittent GnRH stimu lation leading to increased GnRH receptor expression; this is a

central facet of the self-­priming effect of GnRH and an important mechanism by which GnRH action is modulated in different physiologic states. A majority of gonadotropes synthesize and secrete both LH and FSH. A detailed description of the intracellular mechanisms of GnRH action on the gonadotrope is provided in Chapter 2. Briefly, GnRH receptor binding activates the guanosine triphosphate (GTP)-­binding protein Gq/11, leading to an increase in second messengers inositol 1,4,5-­ triphosphate (IP3) and 1,2-­diacylglycerol (DAG). Further intracellular signaling involves increased intracellular calcium and activation of various protein kinase C (PKC) isoforms, mitogen-­activated protein kinases (e.g., extracellular signal–regulated kinase [ERK], c-­Jun NH2-­terminal kinase [JNK], p38), and calcium/calmodulin-­dependent kinase II (Ca/CaMK II). A pathway involving adenylate cyclase, cyclic adenosine monophosphate (cAMP), protein kinase A (PKA), and cAMP response element binding proteins (CREBs) also plays a role. Each gonadotropin consists of two protein subunits, α and β. The 92-­amino acid α-­subunit is common to both LH and FSH, in addition to human chorionic gonadotropin (hCG) and TSH. The β-­subunits for LH (LHβ) and FSH (FSHβ) are 121 and 111 amino acids in length, respectively, and account for the biologic specificity of these two hormones. GnRH stimulates gene expression of LHβ, FSHβ, and α-­subunit; the latter noncovalently dimerizes with either LHβ or FSHβ to form LH or FSH, respectively. Gonadotropin subunits also undergo variable posttranslational modification, primarily glycosylation (addition of oligosaccharide moieties to specific asparagine residues); such modifications appear to facilitate gonadotropin assembly and influence gonadotropin bioactivity and elimination half-­ life.46 The gonadotropins are then packaged into secretory granules for eventual secretion. Although GnRH is the primary stimulus for LH and FSH synthesis and release from a common cell type, concentrations of these two gonadotropins vary differentially throughout ovulatory cycles, with FSH predominance in the early follicular phase and LH predominance in the late follicular phase. This sequential pattern of FSH and LH predominance is important

9

50 45 40 35 30 25 20 15 10 5 0

1 Pulse/ hour

20 15 10 5

1 Pulse/ 3 hours

0

1 Pulse/ hour

5 10 15 20 25 30 35 40 Days

500 450 400 350 300 250 200 150 100 50 0

1 LH FSH (ng/mL)

LH (ng/mL)

CHAPTER 1  Neuroendocrinology of Reproduction

Fig. 1.10 Luteinizing hormone (LH) and follicle-­stimulating hormone (FSH) concentrations in gonadectomized (but sex steroid–replaced) monkeys after arcuate nucleus ablation—a model of isolated GnRH deficiency.  Exogenous GnRH administered in a pulsatile fashion every hour reconstituted LH and FSH secretion. Changing GnRH pulse administration from a relatively high frequency (hourly) to a relatively low frequency (every 3 hours) resulted in decreased LH but increased FSH secretion. (Modified from Wildt L, et al. Frequency and amplitude of gonadotropin-­releasing hormone stimulation and gonadotropin secretion in the rhesus monkey. Endocrinology. 1981;109:376–385.)

for normal follicular maturation, ovarian steroid production, and subsequent ovulation. At least two mechanisms govern differential gonadotropin secretion throughout ovulatory cycles. First, both estradiol and inhibins selectively inhibit FSH release from gonadotropes during the mid-­and late follicular phase in addition to the luteal phase.47,48 Second, different patterns of pulsatile GnRH release differentially affect gonadotropin synthesis and secretion. Specifically, high-­ frequency GnRH pulses favor LH production, whereas low-­ frequency GnRH pulses favor FSH production. For example, studies in ovariectomized, GnRH-­deficient monkeys reveal that a decrease in the frequency of exogenously administered GnRH pulses from one pulse per hour to one pulse every 3 hours results in a 65% increase in plasma FSH, despite a 50% decrease in LH (Fig. 1.10).42 Similar findings have been described in sheep49 and humans.50,51,27 Detailed studies in rats demonstrate that rapid GnRH pulse stimulation favors α-­ subunit and LHβ mRNA expression, whereas slow GnRH pulses favor FSHβ mRNA expression.52 The mechanisms effecting differential LH and FSH expression in response to changes in GnRH pulse frequency are complex53–55 but include variations of GnRH receptor number on the gonadotrope cell surface56 and alterations of gonadotrope activin βB and follistatin expression.57 A pulse of GnRH release stimulates a pulse of LH release on a one-­to-­one basis, and LH (or α-­subunit) pulse patterns, as assessed by frequent sampling of peripheral blood, accurately mirror GnRH pulse patterns in animal studies (Fig. 1.11).40,58 Similarly, exogenous GnRH pulses elicit corresponding LH pulses in GnRH-­deficient patients. Because measurable GnRH is effectively confined to the hypophyseal portal system, which is inaccessible in humans, GnRH pulse frequency is inferred from LH pulse frequency (or α-­subunit pulse frequency59,60) in human studies. Although pulses of GnRH stimulate pulsatile release of FSH, the longer serum half-­life of FSH renders FSH pulses more difficult to identify via frequent sampling of peripheral blood. In addition, although short-­term LH secretion is very closely tied to continued GnRH stimulation, FSH secretion is less acutely dependent on GnRH stimulation.61,62 For example, with GnRH antagonism, the percentage reduction in LH exceeds that of FSH.

GnRH

0

1

2

3

4

5

Hours Fig. 1.11 Close temporal relationship between pulses of luteinizing hormone (LH) (jugular vein) and gonadotropin-­releasing hormone (GnRH) (pituitary portal system) in the sheep model. (Modified from Moenter SM, et al. Dynamics of gonadotropin-­releasing hormone release during a pulse. Endocrinology. 1992;130:503–510.)

Neuronal Inputs Into Gonadotropin-­Releasing Hormone Neurons • Normal pulsatile GnRH secretion is dependent on complex interactions among numerous afferent neuronal inputs, including those expressing kisspeptin, neurokinin B, and dynorphin. • According to current models, kisspeptin stimulates GnRH release, whereas neurokinin B and dynorphin modulate GnRH release primarily by stimulation and suppression, respectively, of kisspeptin release.    The governance of GnRH neurons is highly complex and involves numerous interacting neural systems involving various neurotransmitters and neuromodulators. The neuronal populations upstream of the GnRH neuron play key roles in puberty and are important mediators of sex steroid feedback and the influence of nutritional cues and stress on GnRH secretion. Numerous neurotransmitters appear to be involved in the regulation of GnRH secretion—including dopamine, norepinephrine, glutamate, GABA, and nitric oxide—and the recent discovery of several neuronal populations upstream of the GnRH neuron (e.g., kisspeptin neurons) has markedly enhanced our understanding of reproductive neuroendocrinology.

Kisspeptin Kisspeptin-­secreting neurons appear to be requisite for normal GnRH secretion, serving as a “gatekeeper” of puberty and helping to mediate the effects of sex steroids and metabolic cues on GnRH secretion. Kisspeptin was originally called metastin because of its ability to suppress metastatic spread of human melanomas and breast carcinomas. However, in recognition of its discovery at Pennsylvania State University in Hershey, Pennsylvania, it was later named kisspeptin after Hershey’s chocolate KISSES. Herein we will use the following abbreviations64: KISS1 and Kiss1, the human and nonhuman kisspeptin genes, respectively; KISS1R (Kiss1R) and KISS1R (Kiss1R), the human (nonhuman) kisspeptin receptor genes and gene products, respectively. The KISS1 gene product is a 154–amino acid precursor protein (kisspeptin 1-­ 145). Variable proteolytic modification yields kisspeptins of different lengths: kisspeptin-­54, -­14, -­13, and -­10, with the numbers referring to the amino acid length of bioactive kisspeptin fragments (Fig. 1.12). Importantly, functional native kisspeptins terminal (kisspeptin

10

PART I  The Fundamentals of Reproduction 1 Kp-145, precursor

26

145

SP NH2

15.4 kDa 68

121

Kp-54, Metastin

COOH 5.9 kDa

108

121

Kp-14

1.7 kDa 109

121

Kp-13

1.6 kDa 112

Kp-10

121 1.3 kDa

Fig. 1.12 Schematic of the precursor kisspeptin-­145 and the functional kisspeptin (Kp) fragments, including size and cleavage sites.  Note that all functional Kp fragments maintain amino acids 112 to 121 (red). SP, Signal peptide. (Modified from Roseweir AK, Millar RP. The role of kisspeptin in the control of gonadotrophin secretion. Hum Reprod Update. 2009;15:203–212.)

amino acids 112 to 121), which are important for receptor binding and function. Kisspeptin is the natural ligand of KISS1R—formerly known as the G protein–coupled receptor 54 (GPR54)—a seven– transmembrane domain, G protein–coupled receptor. The importance of the kisspeptin system in reproduction was initially revealed by members of two consanguineous families with loss-­of-­function KISS1R variants leading to pubertal failure and normosmic hypogonadotropic hypogonadism.65,66 Inactivating KISS1 variants leading to pubertal failure and normosmic hypogonadotropic hypogonadism have also been described in four sisters.67 Murine Kiss1R and Kiss1 knockout models exhibit hypogonadotropic hypogonadism with impaired sexual maturation, reduced gonadal size, failure of estrous cyclicity in females, impaired spermatogenesis in males, and infertility.66,68,69 In contrast, gain-­of-­function KISS1R and KISS1 variants may cause precocious puberty.70,71 KISS1R and KISS1 variants neither interrupt GnRH neuron migration to the hypothalamus nor impair GnRH synthesis. Single boluses of kisspeptin markedly stimulate LH release in rodents, sheep, monkeys, and humans. This effect of kisspeptin is mediated by stimulation of GnRH neurons as supported by the following: kisspeptin fibers form synaptic contacts with GnRH neurons,72,73 which is observable in utero74,75; the kisspeptin receptor is expressed by a majority of GnRH neurons76–78; kisspeptin directly depolarizes GnRH neurons79,80; a kisspeptin antagonist inhibits murine GnRH neuron firing rates and reduces pulsatile GnRH release in female pubertal monkeys81; and kisspeptin stimulation of gonadotropin secretion is completely blocked by GnRH antagonists.64,76,82 However, kisspeptin also appears to work indirectly because kisspeptin can increase GABAergic and glutamatergic postsynaptic currents onto GnRH neurons.83 Kisspeptin does not stimulate LH secretion in Kiss1R knockout mice,69,84 suggesting that kisspeptin acts exclusively through its cognate receptor. Moreover, mice with GnRH neuron-­specific Kiss1R knockout exhibit hypogonadotropic hypogonadism and infertility85,86; but in global Kiss1R knockout mice, restoration of GnRH neuron-­ specific Kiss1R expression restores normal reproductive function.85,86 These findings suggest that kisspeptin action at GnRH neurons is critical for reproductive function. In addition to acting upon GnRH neuron cell bodies,87 kisspeptin neurons extensively synapse with GnRH neuron terminals in the external zone of the median eminence,88 where kisspeptin stimulates GnRH release (exocytosis) into the hypothalamic portal system.78,89 Although kisspeptin may have direct effects on gonadotropes, available data suggest that this does not play a major role in kisspeptin’s ability to stimulate gonadotropin secretion. For example, pulsatile GnRH can restore normal reproductive function in patients with loss-­of-­function KISS1R variants.90 In primates (including humans), the majority of the kisspeptin cell bodies reside in the infundibular (arcuate) nucleus.91–93 In con trast, rodents have two primary populations of kisspeptinneurons in the hypothalamus: one in the arcuate nucleus (mediobasal

hypothalamus), and the other in the rostral periventricular area of the third ventricle (RP3V) of the preoptic area, which includes the anteroventral periventricular nucleus (AVPV).94 Of interest, kisspeptin expression in the AVPV is much higher in female compared with male rodents, which appears to reflect organizational effects of sex steroids during early development95,96; and kisspeptin neurons in the AVPV appear to be specifically important for LH surge generation in rodents. Sexual dimorphism of kisspeptin expression has also been described in sheep97 and humans.92 Although two studies suggest that adult women exhibit kisspeptin neurons in the rostral hypothalamus,91,98 it remains unclear whether or not such kisspeptin neurons are homologous to those in the rodent AVPV/RP3V. Kisspeptin’s ability to stimulate LH release in women may vary according to cycle phase or hormonal milieu. For example, although bolus kisspeptin administration consistently increases LH release in women studied in the luteal and preovulatory phases, its effects are less consistent when administered in the early to mid-­follicular phase.99–101 Such differences may reflect the observation in some studies that LH responses to kisspeptin positively correlate with circulating estradiol concentrations.102,103 However, compared with findings in cycling women studied during the follicular phase, acute LH responses to bolus kisspeptin administration appear to be more pronounced in women with functional hypothalamic amenorrhea.104 Moreover, one study suggested that LH responses to bolus kisspeptin are more pronounced in postmenopausal women,105 although another study suggested that 24-­hour kisspeptin infusions do not increase LH release in hypoestrogenic postmenopausal women.103 Kisspeptin and its analogues may hold therapeutic utility in the future.106 Kisspeptin has been investigated in several disorders marked by impaired GnRH secretion and low gonadotropins: functional hypothalamic amenorrhea,104,107,108 hypogonadism associated with obesity and diabetes,109 and hyperprolactinemia.110 Rapid proteolytic degradation of kisspeptin may limit its therapeutic utility, however. Although long-­acting KISS1R agonists are actively being developed, the precise effects of long-­ term KISS1R agonism on gonadotropin release remain unclear; in this regard, desensitization to kisspeptin may represent a practical challenge. Kisspeptin has been evaluated as a trigger for final oocyte maturation and ovulation in women at risk for ovarian hyperstimulation syndrome.111,112 In this scenario, rapid proteolytic degradation may be advantageous as compared to hCG. Finally, when complete gonadal steroid suppression is not required (e.g., endometriosis), KISS1R antagonists may permit partial inhibition of gonadotropin production.

Neurokinin B Neurokinin B—a decapeptide (Asp-Met-His-Asp-Phe-­Phe-­Val-­ ) product of the tachykinin 3 gene (TAC3)—is a member of the tachykinin family, which also includes substance

CHAPTER 1  Neuroendocrinology of Reproduction α-Neoendorphin

Prodynorphin

11

Dynorphin A

1

Dynorphin B Proenkephalin

Peptide F

OctaPeptide

HeptaPeptide

POMC

γ-MSH Leu Enkephalin

α-MSH CLIP ACTH

β-MSH γ-LPH

β-Endorphin β-LPH

Met Enkephalin

Fig. 1.13 Schematic of endogenous opiate precursors.  ACTH, Adrenocorticotropic hormone; CLIP, corticotropin-­like intermediate lobe peptide; LPH, lipotropin; MSH, melanocyte-­stimulating hormone; POMC, proopiomelanocortin. (Modified from Akil H, et al. Endogenous opioids: overview and current issues. Drug Alcohol Depend. 1998;51:127–140.)

P (SP) and neurokinin A (NKA), which are products of the TAC1 gene. There are several neurokinin receptors (NK1R, NK2R, NK3R), and although NKB can produce some agonism at NK1R and NK2R, NKB binds preferentially to and acts primarily via its cognate receptor NK3R (encoded by the TACR3 gene).113 Studies of patients with idiopathic hypogonadotropic hypogonadism from consanguineous families revealed that homozygous loss-­of-­ function variants of either TAC3 or TACR3 can cause pubertal failure and severe hypogonadotropic hypogonadism, highlighting the importance of NKB in human reproduction.114,115 The role of NKB in central reproductive function is complex and appears to vary according to species, sex, and sex steroid milieu.116,117 The selective NK3R agonist senktide can stimulate LH secretion—albeit not as potently as kisspeptin—in rats,118 sheep,119 and monkeys.120 Such stimulation of LH secretion by NKB is mediated by GnRH secretion, and GnRH receptor antagonism abolishes LH responses to senktide in the monkey.120 Although it remains unclear to what degree NKB may have direct actions on GnRH neurons,121–123 a number of observations suggest that NKB primarily influences pulsatile GnRH secretion indirectly by stimulating kisspeptin release. For example, kisspeptin neurons express NK3R, and senktide increases kisspeptin neuronal activity.118 LH responses to senktide are either absent or markedly reduced in Kiss1R knockout mice,124 in the presence of Kiss1R antagonism,125 or after Kiss1R desensitization.126 Moreover, continuous kisspeptin infusion can restore pulsatile LH secretion in patients with loss-­of-­function variants of TAC3 or TACR3.127 In contrast to Kiss1 and Kiss1R knockout mice, Tacr3 knockout mice remain fertile, although they can demonstrate reproductive defects.128,129 This apparent discordance may reflect redundancy in the roles of SP-­NK1R, NKA-­NK2R, and NKB-­NK3R in rodents. For example, SP, NKA, and NKB can all activate kisspeptin neurons in mice,130,131 and antagonism of NK1R, NK2R, and NK3R is required to block NKB activation of murine kisspeptin neurons.130 In contrast, less redundancy is evident in ruminants and primates.132 Regarding the therapeutic potential of NKB analogues, two studies in women with polycystic ovary syndrome (PCOS, a

disorder marked by persistently high GnRH pulse frequency, LH excess, and hyperandrogenemia) suggested that selective NK3R antagonism for 7 days reduces LH (GnRH) pulse frequency, LH area under the curve, serum LH concentrations, basal (non-­ pulsatile) LH secretion, and total testosterone concentrations, with essentially no change in estradiol concentrations.133,134 Short-­term studies also suggest that selective NK3R antagonists may be useful in disorders requiring only partial reductions in gonadotropins and gonadal steroids (e.g., endometriosis).135–138 Such potential uses require further study, however, as the impact of chronic NK3R antagonism has not yet been assessed in premenopausal women. Also of interest, accumulating data suggests that increased NKB signaling plays a role in the vasomotor symptoms associated with estrogen deficiency,139 and early clinical trials suggest that NK3R antagonism ameliorates menopausal hot flashes.140–142

Endogenous Opioid Peptides Endogenous opioid peptides (EOPs), which include endorphins, enkephalins, and dynorphins, participate in myriad processes such as motor activity, cognitive functions, water and food intake, and regulation of neuroendocrine function.143 Most active EOPs share a common sequence (Tyr-­ Gly-­ Gly-­ Phe-­ [Met or Leu]) at the amino-­terminal, although endorphins, enkephalins, and dynorphins are derived from different precursor proteins that undergo regulated posttranslational processing (Fig. 1.13).144 Endorphins such as β-­endorphin are products of the precursor protein POMC. POMC can be preferentially processed to produce ACTH and β-­ lipotropin, as occurs in corticotropes (adenohypophysis) under the control of CRH. However, in the hypothalamus, POMC processing primarily yields β-­endorphin and α-­melanocyte-­stimulating hormone. Hypothalamic β-­ endorphin participates in the regulation of reproduction, temperature, and cardiovascular and respiratory functions, and acts opioid receptors. Enkephalins are derived from proenkephalin, and their primary functions appear to relate to

12

PART I  The Fundamentals of Reproduction

autonomic nervous system modulation, mainly via δ-­receptor activation. Dynorphins are products of the precursor prodynorphin and act chiefly at κ-­opioid receptors (KORs). Importantly, although β-­endorphin, enkephalins, and dynorphins act primarily via μ-­, δ-­, and κ-­opioid receptors, respectively, each can act as agonists at more than one receptor subtype. Numerous studies provide evidence that hypothalamic opiates partly mediate sex steroid negative feedback on GnRH release. For example, GnRH neurons express few, if any, progesterone receptors, whereas β-­endorphin concentrations increase in hypophyseal blood during the luteal phase in monkeys (when progesterone suppresses GnRH pulse frequency).145,146 Moreover, naloxone and naltrexone (opiate receptor antagonists acting primarily at μ-­ and κ-­opioid receptors) increase LH (GnRH) pulse frequency when administered to luteal phase women147 or progestin-­ treated postmenopausal women.148 Similarly, morphine suppresses GnRH secretion from mediobasal hypothalami isolated from fetal and adult humans—an effect that is reversed by naloxone—and chronic high-­dose opiate administration can cause hypogonadotropic hypogonadism by suppressing GnRH and LH secretion.143,149 Several animal studies implicate dynorphin as a principal mediator of progesterone negative feedback on GnRH pulse frequency in females.87 Progesterone treatment in ewes increases dynorphin A concentrations in third ventricle cerebrospinal fluid,150 and central infusion of dynorphin in goats reduces volleys of multiple-­unit activity in the mediobasal hypothalamus and reduces LH pulses.151 In luteal phase ewes, specific κ-­opioid receptor antagonists—but not antagonists to δ-­ or μ-­opioid receptors—reverse progesterone inhibition of LH secretion and LH pulse frequency when locally administered into the mediobasal hypothalamus.152 Dynorphin may exert its mediating actions directly on GnRH neurons: for example, dynorphin neurons in the arcuate nucleus colocalize with progesterone receptors in ewes,153 and dynorphin-­containing varicosities are closely associated with GnRH neuron cell bodies in the mediobasal hypothalamus.152,154 As described in more detail below, dynorphin also appears to influence kisspeptin release.155 In addition, other EOPs (e.g., β-­endorphin) in other hypothalamic areas may also be involved in the control of GnRH release. For example, in the aforementioned study,152 κ-­ and μ-­receptor antagonists locally administered into the preoptic area increased LH and LH pulse frequency.

Kisspeptin, Neurokinin B, Dynorphin (KNDy) Neurons In the arcuate nucleus, kisspeptin, NKB, and dynorphin are frequently coexpressed in the same neuron. For example, kisspeptin neurons in the arcuate nucleus have been found to coexpress NKB and dynorphin in rodents,156,157 goats,151 and sheep.158 For convenience, and as a nod to kisspeptin (namesake of Hershey’s chocolate KISSES), such neurons are often called KNDy neurons (Kisspeptin, Neurokinin B, Dynorphin; pronounced candy).97 KNDy neurons in the arcuate nucleus form an extensively interconnected network.156,159,160 KNDy axons also project to the internal zone of the median eminence where they are in close proximity to GnRH fibers.121,161 As with kisspeptin neurons, KNDy neuron neuroanatomy exhibits sexual dimorphism, possibly related to perinatal sex steroid exposure.97 In addition, robust experimental data in rodents and ruminants suggest that KNDy neurons are intimately involved with sex steroid feedback on GnRH secretion, and general consensus holds that a subpopulation of KNDy neurons represents a fundamental component of the GnRH pulse generator in these species.87,162 Corresponding data in humans are limited. In one autopsy study, 77% of kisspeptin cell bodies (and 56% of kisspeptin axon fibers) in the infundibular nucleus coexpressed preproNKB, and 95% of preproNKB-

kisspeptin.92 However, the degree of colocalization in humans appears to differ according to sex and age. For example, one autopsy study suggested that only 10% and 26% of kisspeptin-­ containing afferent contacts onto GnRH neurons coexpressed preproNKB in older men and women, respectively163; another autopsy study in young men suggested that 75% of infundibular kisspeptin-­containing cell bodies also contained NKB, 33% of NKB-­containing cell bodies also contained kisspeptin, and colocalization with dynorphin was uncommon.164 Although these small studies suggested limited colocalization in humans, it is unclear to what degree postmortem degradation may have influenced these findings. Regardless, it remains well accepted that kisspeptin, NKB, and EOPs (e.g., dynorphin)—released from neurons that do or do not colocalize with the other peptides— substantially influence GnRH neuronal function in humans.

Gonadotropin-­Inhibitory Hormone and RFamide-­Related Peptides The roles of gonadotropin-­inhibitory hormone (GnIH) and its mammalian orthologues, RFamide-­ related peptides (RFRPs), in the central control of reproduction have been recently reviewed.165,166 Briefly, RFRP-­immunoreactive cells have been identified in hypothalami of a number of species, including RFRP-­ 1 and RFRP-­ 3 in humans.167 RFRP-­immunoreactive fibers project to GnRH neurons in the median eminence in rhesus macaques and humans,167,168 in addition to a subset of arcuate kisspeptin neurons in mice.169 RFRP-­3 can reduce GnRH neuronal firing rates in mice170; RFRP-­3 inhibits pituitary gonadotropin release from cultured ovine pituitary cells171; and intravenous RFRP-­3 administration suppresses LH pulse amplitude in ovariectomized ewes.172 One study revealed reduced RFRP expression in the preovulatory period in ewes, suggesting a reciprocal relationship with GnRH release, and infusion of GnIH blocked the estrogen-­induced LH surge.173 In contrast, RFRP precursor expression did not decrease in the late follicular phase in a study of rhesus macaques, suggesting the possibility of species differences in this regard.174 GnIH and RFRPs have been implicated in the regulation of gonadal function (decrease), food intake (increase), and sexual motivation (decrease), in addition to mediating the inhibitory influence of stress on reproduction.165,166 Although a growing body of data suggests that RFRPs are important factors controlling GnRH and gonadotropin secretion in a number of mammalian species, an understanding of their precise role(s) in humans awaits further investigation.

Gonadotropin-­Releasing Hormone Pulse Generator • Discrete, intermittent bursts of coordinated GnRH neuron activity lead to pulsatile release of GnRH into the hypophyseal portal system. • Although pulsatility is an intrinsic property of GnRH neurons, afferent inputs (e.g., neurons expressing kisspeptin, NKB, and/ or dynorphin) are required for normal GnRH pulse generation and appear to represent integral components of the GnRH pulse generator.    As described previously, intermittent GnRH receptor stimulation is an absolute requirement for physiologic maintenance of gonadotropin secretion. Although the precise basis of pulsatile GnRH release remains unclear, a number of observations strongly support the concept that neuronal systems within the mediobasal hypothalamus effect pulsatile release of GnRH into the hypophyseal portal system. In animal models, volleys of multiple unit electrical activity (i.e., detection of activity in multiple neurons near an electrode) in the area of the mediobasal hypothalamus coincide with the initiation of LH pulses (Fig. 1.14).175,176 Similarly, electrical stimulation via electrodes placed in the mediobasal

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Time (min) Fig. 1.14 Temporal association between volleys of multiple unit activity (MUA) in the hypothalamus and luteinizing hormone (LH) pulses (green) detected in peripheral blood in an ovariectomized monkey. (Modified from Knobil E. The electrophysiology of the GnRH pulse generator in the rhesus monkey. J Steroid Biochem. 1989;33:669–671.)

hypothalamus stimulates GnRH release into the hypophyseal portal system in monkeys.177 Mediobasal hypothalami isolated from both fetal (20 to 23 weeks’ gestation) and adult humans release GnRH in discrete pulses with a frequency approximating one pulse per 60 to 100 minutes,149 and mediobasal hypothalami separated from the remainder of the brain can maintain pulsatile LH secretion in monkeys.178 These data suggest that the mediobasal hypothalamus houses all requisite components for GnRH pulse generation (i.e., the GnRH pulse generator) and that pulsatile GnRH release does not require innervation from outside of the mediobasal hypothalamus. Nonetheless, mechanisms underlying episodic GnRH pulse generation, and what neuroanatomic components constitute the GnRH pulse generator, remain uncertain. Several studies suggest that pulsatility is an intrinsic property of GnRH neurons. For example, pulsatile GnRH release is exhibited by immortalized GnRH-­secreting neurons179,180 and by cultured GnRH neurons obtained from fetal rats, sheep, and monkeys.181–183 If GnRH pulse generation reflects an intrinsic property of GnRH neurons, then coordination of GnRH release could be facilitated by cell-­ to-­ cell interconnections among GnRH neurons.14,184 It is well accepted that afferent inputs (e.g., kisspeptin neurons) are important for normal GnRH secretion, and accumulating research supports the hypothesis that kisspeptin (KNDy) neurons represent a fundamental component of the GnRH pulse generator, in essence orchestrating coordinated GnRH neuronal activity and GnRH secretion accordingly.87,162,185 As described previously, LH pulses are temporally associated with volleys of multiunit activity in the arcuate nucleus, which contains both GnRH and kisspeptin (KNDy) neurons.186 In addition, kisspeptin release at the median eminence appears to be pulsatile: although kisspeptin pulses were not clearly coincident with peripheral LH pulses in ovariectomized ewes,187 kisspeptin pulses corresponded to GnRH pulses 75% of the time in midpubertal rhesus monkeys.188 Moreover, a GnRH neuron cell culture study suggests that pulsatile kisspeptin administration entrains synchronous cycles of GnRH gene transcription and pulsatile GnRH secretion.189 Similarly, a recent in vivo mouse study indicates that episodic increases in arcuate kisspeptin neuron activity correlate very highly with LH pulse generation, and brief optogenetic activation of arcuate kisspeptin neurons generated LH pulses.190 However, work in sheep suggests that additional elements (e.g.,

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upstream glutamate-­secreting neurons) may also be important in this regard.191 Therefore, although kisspeptin neurons appear to be an important mediator of GnRH pulse secretion, the fundamental nature of the GnRH pulse generator remains unclear. Human studies also imply that kisspeptin plays a role GnRH pulse generation. For example, in men, continuous intravenous infusion of a relatively low dose of kisspeptin can increase LH pulse frequency192; and a single injection of kisspeptin may reset the GnRH pacemaker.193 (In the latter study, the interval between the kisspeptin-­induced LH pulse and the immediately preceding endogenous LH pulse was variable but on average shorter than the normal LH interpulse interval; in contrast, the interval between the kisspeptin-­induced LH pulse and the subsequent endogenous LH pulse was similar to normal interpulse intervals [approximately 2 hours], suggesting that kisspeptin administration reset the hypothalamic GnRH clock.) Parallel results in women are mixed: although bolus kisspeptin administration did not appear to reset the GnRH pacemaker in women,101 single-­ dose subcutaneous kisspeptin administration during the follicular phase has been reported to increase LH pulse frequency.194 NKB and dynorphin may also play important roles in the coordination of pulsatile GnRH release. This notion is consistent with a number of experimental observations. For example, KNDy neurons exhibit both NK3R and κ-­opioid receptors87,195; murine kisspeptin neuron firing rates are increased by NK3R agonists and reduced by κ-­opioid receptor agonists130,196—effects that appear to be modulated by gonadal steroids.196,197 In addition, central administration of dynorphin in goats inhibits both multiple unit activity (MUA) volleys in the mediobasal hypothalamus and pulsatile LH release, whereas NKB provokes MUA volleys.151 Studies using microimplants in the arcuate nucleus of ewes revealed consistent findings: LH pulse frequency was decreased by an NK3R antagonist, whereas LH pulse frequency was increased by either NKB or a κ-­opioid receptor antagonist.198 Fig. 1.15 depicts a working model proposed by Moore et al., primarily based on experiments performed in sheep.155 According to this model, KNDy neurons signal to other KNDy neurons— and perhaps to other neurons within the arcuate nucleus—with NKB release stimulating kisspeptin secretion, which in turn initiates GnRH pulse secretion. Subsequent dynorphin release then inhibits kisspeptin secretion, effecting GnRH pulse termination. Importantly, this KNDy hypothesis of pulsatile GnRH secretion is largely based on detailed studies in rodents and ruminants (sheep, goats), and a full understanding of how these data relate to humans awaits further research.132,162 Some data suggest that kisspeptin may not be required for GnRH pulse generation. In particular, frequent sampling studies reveal that humans with loss-­ of-­ function KISSR variants demonstrate pulsatile LH release, albeit at low amplitude.66,199 Similarly, a study suggested that puberty occurs and fertility is preserved in female mice with either (1) congenital absence of kisspeptin neurons or (2) congenital absence of neurons expressing Kiss1R.200 When taken as a whole, available data imply that kisspeptin action may not be an absolute requirement for pulsatile GnRH secretion, but it is clear that kisspeptin is required for normal GnRH pulse secretion and normally exerts a profound influence on GnRH pulse generation.

Gonadotropin-­Releasing Hormone Secretion During Development and in Adulthood • Gonadotropin secretion is robust during fetal development and early infancy but quiescent during childhood; puberty represents the reemergence and amplification of gonadotropin secretion, which stimulates gametogenesis, gonadal sex steroid secretion, and the physical manifestations of puberty. • GnRH pulse frequency changes across the normal menstrual cycle, being highest in the late follicular phase and lowest in the luteal

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Fig. 1.15 Working model regarding how KNDy neurons may participate in gonadotropin-­releasing hormone (GnRH) pulse generation and termination.  By this model, neurokinin B (NKB; green) stimulates and dynorphin (Dyn; red) suppresses kisspeptin (Kp; blue) release, with kisspeptin stimulating GnRH release from GnRH neurons (gray). The onset of a GnRH pulse is triggered by an initial increase in NKB release, which increases kisspeptin output. Kisspeptin may also stimulate interneurons (orange) that support/strengthen NKB stimulation of KNDy neurons. After a short period of time, an increase in DYN release from KNDy neurons suppresses kisspeptin release from KNDy neurons in addition to acting directly on GnRH neurons to inhibit GnRH release. Dynorphin may also inhibit interneurons (orange) that support and strengthen kisspeptin release from KNDy neurons. Note that the neuronal process and terminal color denotes the active neuropeptide at the relevant synapse; it does not indicate selective neuropeptide transport through the neuronal process. The dashed circle represents the arcuate nucleus (ARC). RDyn, kappa opioid receptor; RKp, kisspeptin receptor; RNKB, neurokinin 3 receptor. (Modified from Moore AM, et al. KNDy cells revisited. Endocrinology. 2018;159:3219–3234.)

phase; these day-­to-­day changes primarily reflect the imposition or removal of progesterone negative feedback, and they contribute to the normal cyclic patterns of LH and FSH secretion. Men demonstrate consistent day-­to-­day GnRH pulse patterns, with • GnRH pulse frequency approximating one pulse every 2 hours.

Physiologic Development of Reproductive Neuroendocrine Function Patterns of GnRH secretion change markedly across human development. Reproductive neuroendocrine events throughout early maturation, including both before and during the establishment of reproductive competence, are discussed in detail in Chapter 18. Briefly, GnRH and gonadotropin secretion is robust in utero, peaking in midgestation. In males, gonadotropin secretion markedly stimulates testicular androgen secretion, which is important for normal genital differentiation. The gestational increase in sex steroid (e.g., estradiol) production from the fetoplacental unit provides negative feedback to limit fetal GnRH and gonadotropin secretion. Birth is followed by a marked but transient (3-­to 9-­month) increase in GnRH and gonadotropin secretion (the “minipuberty of infancy”), perhaps related to the withdrawal of fetoplacental sex steroids. A marked sex difference of gonadotropin release is evident at this time, with LH concentrations being higher in males and FSH levels higher in females. The possibility that kisspeptin is important for the minipuberty of infancy is suggested by a patient with a compound heterozygote loss-­of-­function KISSR variant, who had micropenis, undescended testes, and undetectable serum gonadotropins at 2 months of age—a time usually marked by robust gonadotropin secretion.201

By late infancy or early childhood (earlier in boys than in girls), GnRH and gonadotropin secretion markedly decreases, leading to a hypogonadotropic phase of childhood marked by low sex steroid concentrations—the juvenile pause. Studies of gonadotropin secretion in children reveal low LH and FSH concentrations, a high FSH-­to-­LH ratio, and low LH pulse amplitude and frequency.202 Mechanisms accounting for low GnRH secretion during this time appear to include inhibition of the GnRH pulse generator (neurobiologic brake) by higher-­order neuronal systems (e.g., involving GABA-­and NPY-­secreting neurons) and a developmental removal of stimulation (e.g., involving neurons secreting glutamate and norepinephrine). Near the close of the first decade, a marked nocturnal amplification of pulsatile LH secretion indicates the neuroendocrine initiation of puberty. A majority of studies suggest that early pubertal subjects demonstrate sleep-­entrained increases in LH (GnRH) pulse frequency and amplitude.203 Gonadotropin concentrations rise across puberty,204,205 stimulating gametogenesis, gonadal sex steroid secretion, and the development of secondary sexual characteristics. Mechanisms underlying puberty are poorly understood, but they likely reflect developmental remodeling of inhibitory and stimulatory neural circuits in the hypothalamus. For example, puberty has been associated with reductions of GABAergic inhibitory neurotransmission and an increase in excitatory neurotransmitters such as glutamate. Kisspeptin and NKB also appear to play critically important roles in human puberty because inactivating variants of KISS1, KISS1R, TAC3, or TACR3 result in pubertal failure. Conversely, central precocious puberty has been associated with gain-­of-­function KISS1R variants70 and KISS1 variants that may impair kisspeptin degradation.71 In addition, loss-of-function variants in the maternally imprinted genes

CHAPTER 1  Neuroendocrinology of Reproduction

15

MKRN3 (encoding makorin ring finger protein 3) and DLK1 (encoding delta-­like non-­canonical Notch ligand 1) have been discovered as causes of central precocious puberty,206–210 suggesting that MKRN3 and DLK1 contribute to the neurobiologic brake. In addition to these transsynaptic mechanisms, neuroglial cells may contribute to the pubertal reactivation of GnRH secretion (e.g., by secretion of growth factors).2

• In men, GnRH secretion is restrained by testosterone and dihydrotestosterone (DHT) in addition to estradiol, a product of testosterone aromatization, but estradiol is the primary mediator of testosterone negative feedback at pituitary gonadotropes. • Afferent neuronal pathways (e.g., kisspeptin and dynorphin neurons) are key mediators of sex steroid negative feedback on GnRH secretion.   

Patterns of Pulsatile Gonadotropin-­Releasing Hormone Secretion in Adults

After puberty, gonadal hormones continually relay information about the state of gonadal function to the hypothalamic-­ pituitary axis. Hypothalamic areas involved with the regulation of GnRH secretion (and pituitary gonadotropes) express receptors for estrogen, progesterone, and androgen, and sex steroid feedback plays a predominant role in the physiologic modification of GnRH and gonadotropin secretion. These steroid feedback signals can thus alter gonadotropin feedforward signals to the gonads by influencing GnRH secretion, modulating pituitary (gonadotrope) responses to GnRH, or both. Under normal circumstances, these regulatory feedback loops maintain appropriate gonadal function. The negative feedback actions of pharmacological doses of sex steroids (e.g., combined oral contraceptives) suppress gonadotropins and can be used for temporary contraception in women. Similar strategies are being developed for men.221

Human studies using frequent blood sampling and formal pulse analysis have documented significant changes of LH (and by inference GnRH) pulse frequency throughout ovulatory cycles. Briefly, average LH (GnRH) pulse frequency is around one pulse every 90 minutes in the early follicular phase, and this gradually increases to approximately one pulse per hour by the late follicular phase. Although monkey studies suggest that GnRH pulse frequency slows during the mid-­cycle surge,211 human studies suggest no change in either LH or α-­subunit pulse frequency at mid-­ cycle.212,213 LH pulse frequency decreases markedly during the luteal phase, approximating one pulse every 3 to 8 hours. These day-­to-­day changes of GnRH pulse frequency appear to be important for normal hormonal changes across ovulatory cycles.214,215 In adult humans and nonhuman primates, GnRH pulses occur approximately once per hour in the (near) absence of sex steroid negative feedback (e.g., after surgical or natural menopause).216–218 Similarly, the isolated human mediobasal hypothalamus secretes GnRH pulses every 60 to 100 minutes,149 and LH pulses do not appear to exceed a once-­hourly frequency during any phase of the cycle.217,219 These findings have contributed to the concept that a pulse frequency of approximately one per hour (a circhoral frequency) may be an inherent characteristic of the adult GnRH pulse generator and that day-­to-­day changes of GnRH pulse frequency in women reflect the imposition or removal of sex steroid (primarily progesterone) negative feedback. LH pulse amplitude also changes across the menstrual cycle. LH pulse amplitude decreases slightly across the follicular phase, but it is greatly amplified at mid-­cycle (i.e., during the LH surge). During the luteal phase, LH pulse amplitude is variable, but in general, it is approximately twofold higher than that of the follicular phase. It is important to note that the amplitude of LH pulses can be modulated centrally via changes of GnRH released per pulse, at the pituitary gonadotrope via changes of gonadotrope responsiveness to GnRH, or both. Also of interest, LH pulse amplitude varies inversely with the preceding LH interpulse interval, which contributes to higher LH pulse amplitudes during the luteal phase.220 In women, dynamic changes of gonadotropin secretion are required to achieve follicular development, ovulation, and preparation for possible pregnancy. In contrast, young men demonstrate consistent daily patterns of GnRH and gonadotropin secretion, with GnRH pulse frequency approximating one pulse every 2 hours. This achieves continuous spermatogenesis, and healthy men are prepared for fertilization at all times. In addition, day-­ to-­ day testosterone secretion remains relatively constant, although testosterone concentrations exhibit diurnal changes with peaks in the morning.

Feedback Regulation of Gonadotropin-­Releasing Hormone and Gonadotropin Secretion • Throughout most of the cycle in women, estradiol restrains GnRH pulse amplitude and gonadotropin secretion (whereas progesterone restrains GnRH pulse frequency), but high estradiol concentrations at mid-­cycle exert positive feedback on pituitary gonadotropes, provoking marked gonadotropin release (the gonadotropin surge).

Negative Feedback Regulation of Gonadotropin-­Releasing Hormone and Gonadotropin Secretion in Women In women, estradiol concentrations correspond to follicular development during the follicular phase and corpus luteum function in the luteal phase. When concentrations are relatively low (i.e., excluding preovulatory concentrations), estrogens restrain gonadotropin secretion. The absence of such feedback accounts for markedly increased gonadotropin secretion in estrogen-­ deficient states such as menopause222 and aromatase deficiency223 (i.e., in open-­ loop conditions). The negative feedback effects of estradiol appear to be mediated primarily at the hypothalamus.224 GnRH release (by direct measurement) is increased in ovariectomized sheep and monkeys, and this is reversed with estrogen replacement.225,226 In human studies, GnRH release can be estimated using GnRH antagonists, with the premise that the degree of LH suppression after incomplete GnRH antagonism is inversely related to endogenous GnRH secretion. In postmenopausal women, the percent reduction in LH concentrations after incomplete GnRH receptor blockade is increased by estradiol replacement.227 Overall, studies indicate that estradiol reduces GnRH pulse amplitude but not GnRH pulse frequency.226,228 Although one study in ovariectomized monkeys suggested that estradiol reduces the frequency of both hypothalamic multiple unit electrical activity and LH pulses,229 available studies in postmenopausal women imply that estrogen replacement primarily reduces LH pulse amplitude rather than LH pulse frequency.218,230 In addition, LH pulse frequency is maximal at one pulse per hour during the late follicular phase in women, when estradiol concentrations are relatively high. Estrogens may also decrease pituitary LH responses to GnRH, although available data are mixed. For example, although estradiol acutely reduces LH release in GnRH-­ deficient monkeys and sheep receiving fixed-­dose exogenous pulsatile GnRH,231,232 relatively low (i.e., not preovulatory) doses of estradiol do not markedly influence LH release in GnRH-­deficient women receiving fixed-­dose exogenous pulsatile GnRH.47 Interestingly, initial reductions of LH release with higher-­dose estradiol may be followed by increased LH release233; this biphasic pattern presumably reflects initial negative feedback and later positive feedback.

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Progesterone is the primary negative feedback regulator of day-­to-­day GnRH pulse frequency in women. LH pulse frequency slows in tandem with early luteal increases in serum progesterone concentrations (from the corpus luteum),219 and LH pulse frequency is inversely correlated with progesterone (but not estradiol) concentrations during the luteal-­follicular transition.234 Moreover, administration of progesterone to women during the follicular phase, when progesterone concentrations are usually low, slows LH pulse frequency.235 Similarly, progesterone plus low-­dose estradiol, but not estradiol alone, slows LH pulse frequency in postmenopausal women.218,230 Importantly, the ability of progesterone to slow GnRH pulses appears to require the permissive presence of estradiol,236,237 which likely reflects the ability of estrogen to increase hypothalamic progesterone receptor expression.238,239 In contrast, progesterone inhibition of GnRH pulse frequency appears to be antagonized by androgens. For example, androgens increase GnRH neuronal firing rates in the mouse model240; and in hyperandrogenic women with PCOS, high GnRH pulse frequencies are relatively resistant to negative feedback suppression by progesterone and estradiol,241 a defect that can be reversed by androgen receptor blockade.242 These findings may be a consequence of androgen-­mediated reductions of hypothalamic progesterone receptor expression.243

Positive Feedback and the Mid-­Cycle Gonadotropin Surge Most examples of endocrine feedback regulation involve negative feedback loops. However, the ovulatory menstrual cycle is unique in that it also depends on the positive feedback effects of sex steroids on the hypothalamic-­pituitary axis. Specifically, high estradiol concentrations from the dominant ovarian follicle provoke a marked increase in gonadotropin release—a mid-­cycle (or preovulatory) gonadotropin surge. In effect, estradiol from the preovulatory follicle signals to the hypothalamic-­pituitary axis that follicular development is adequate for ovulation. Estradiol positive feedback appears to be related to both achieved estradiol concentrations and the duration of estradiol elevation, as demonstrated in both monkeys244 and women.245 Although the mid-­cycle surge is characterized by a marked discharge of both gonadotropins, the increase in LH release exceeds that of FSH, with blood concentrations increasing approximately tenfold versus fourfold, respectively; thus it is often called the LH surge. The mid-­cycle gonadotropin surge uniformly involves positive feedback at the pituitary, markedly increasing gonadotrope responsiveness to GnRH stimulation.246,247 However, the degree to which the gonadotropin surge depends on positive feedback at the hypothalamus appears to be species dependent. GnRH secretion is augmented during the LH surge in rats248 and sheep249; such GnRH surges appear to be physiologically important in these species. Similarly, GnRH release appears to increase in response to estradiol positive feedback in female monkeys.250,251 However, LH surges can be induced by high estradiol concentrations in GnRH-­deficient monkeys receiving constant dose (exogenous) GnRH delivered as once-­hourly pulses,252 implying that a GnRH surge is not essential for LH surge generation in these animals. One study using incomplete GnRH receptor antagonism to estimate GnRH secretion in women suggested that GnRH secretion may be reduced at mid-­cycle compared with the late follicular and early luteal phases.253 Similarly, pulsatile administration of constant-­ dose exogenous GnRH produces LH surges in GnRH-­deficient women247; LH surges can occur in these women even when pulsatile GnRH doses are reduced at mid-­cycle.254 In addition, although pituitary metabolic activity (by positron emission tomography) increases in women at mid-­cycle, hypothalamic metabolic activity does not.255 Thus, although continued GnRH stimulation plays a critically important permissive role in LH surge generation in women (e.g., the surge can be prevented with

GnRH receptor antagonists256,257), available data do not suggest that the gonadotropin surge is accompanied by increased GnRH release in women. Progesterone increases pituitary gonadotropin responses to GnRH,258,259 and the progesterone-­receptor antagonist mifepristone decreases mean LH, LH pulse amplitude, and LH responses to exogenous GnRH when given during the luteal phase.260,261 Nonetheless, in ovariectomized but estradiol-­replaced women, progesterone alone (i.e., without high-­dose estradiol) is unable to induce gonadotropin surges. Indeed, progesterone can block LH surge generation when administered before high-­dose estradiol.262 However, progesterone augments gonadotropin secretion in the setting of preovulatory estradiol concentrations.245,263 Although estradiol alone can provoke an LH surge, the late follicular rise in progesterone (which begins approximately 12 hours before the LH surge)245,264 may be important for the full expression of the mid-­cycle gonadotropin surge. For example, progesterone may increase the duration of the surge,245 and the progesterone-­ receptor antagonist mifepristone can delay the surge.265 Some studies suggest that progesterone may be important for the increase in FSH at mid-­cycle,262,263 whereas others suggest that estradiol alone is sufficient to produce a normal FSH surge.245,266 Of interest, although humans demonstrate sexual dimorphism of hypothalamic neuronal populations (e.g., kisspeptin), the circuitry required for LH surge-­like activity appears to be present in male primates. For example, in male monkeys orchiectomized after puberty, estradiol administration can induce LH surges,267 and ovarian transplants can induce LH surges and other neuroendocrine changes that maintain cyclic function of the transplanted ovary.268 Estradiol and progesterone positive feedback can be experimentally induced in adult men,269,270 but this is not a normal occurrence in male physiology.

Negative Feedback Regulation of Gonadotropin-­Releasing Hormone and Gonadotropin Secretion in Men In contrast to cyclic changes in women, normal men demonstrate a relatively constant average LH pulse frequency of approximately one pulse every 120 minutes, related to consistent tonic inhibition by relatively stable day-­to-­day sex steroid concentrations. In male monkeys, bilateral orchiectomy increases mean LH, LH pulse frequency, and LH pulse amplitude—effects that are prevented by physiologic testosterone replacement.271 Similarly, testosterone-­deficient men (e.g., related to either primary testicular failure or inhibition of steroidogenesis with ketoconazole) exhibit elevated mean LH, LH pulse frequency, and LH pulse amplitude—changes that are at least partially reversed by testosterone replacement.272–274 The importance of the androgen receptor in mediating testosterone negative feedback of LH secretion is suggested by elevated LH concentrations in the setting of androgen insensitivity275 and androgen-­receptor blockade.276,277 Moreover, mean LH is reduced in men by administration of DHT, a potent androgen that cannot be aromatized to estradiol.278–280 However, a portion of synthesized testosterone is aromatized to estradiol, either in testicular Leydig cells or in nongonadal tissues, and estrogens can exert negative feedback actions at the hypothalamic-­pituitary axis. For example, estradiol administration reduces LH secretion in normal and agonadal men.276,278,281,282 Taken together, these findings suggest that both androgens and estrogens exert negative feedback actions at the hypothalamic-­pituitary axis. Many studies suggest that sex steroid negative feedback on the GnRH pulse generator is partly mediated by the androgen receptor. For example, DHT can reduce LH pulse frequency in men,280 and some studies,276,277 but not all,283 suggest that androgen receptor blockade increases LH pulse frequency in men. Similarly, the marked increase in LH pulse frequency with

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high-­dose ketoconazole (which inhibits both testicular/adrenal steroidogenesis and aromatase activity) is completely reversed by testosterone replacement despite low estradiol concentrations, whereas LH pulse frequency is only partly normalized with estradiol add-­back alone (with persistently low testosterone concentrations).274 On the other hand, aromatase inhibitors284 and antiestrogens281,285–287 increase LH pulse frequency in normal men, and estrogen administration to men with aromatase deficiency reduces LH pulse frequency.288 Overall, these data suggest that both androgens and estrogens mediate negative feedback at the GnRH pulse generator. In contrast to dual steroid (androgen, estrogen) restraint of pulsatile GnRH pulse secretion, a number of studies suggest that estradiol is a primary mediator of negative feedback at pituitary gonadotropes in men.274,278,282,289 For example, androgen-­ receptor blockade does not alter LH responses to exogenous GnRH in some studies,277,283 and aromatase inhibition appears to prevent testosterone’s ability to reduce GnRH-­stimulated LH and FSH secretion.273 Perhaps the most compelling studies were performed in GnRH-­deficient men receiving pulsatile exogenous GnRH in constant doses. Under this GnRH-­clamp paradigm, testosterone alone and estradiol alone reduced LH and FSH concentrations, but DHT alone did not.289 In another GnRH-­clamp study, LH pulse amplitude and mean LH increased after high-­ dose ketoconazole, and LH parameters were normalized with estradiol, but not testosterone, add-­back.274 Overall, these study results imply that estradiol is the primary mediator of testosterone negative feedback at the pituitary gonadotrope.

Kisspeptin and KNDy Neurons as Mediators of Sex Steroid Feedback on Gonadotropin-­Releasing Hormone Secretion Evidence in rodents, sheep, and monkeys suggests that sex steroid feedback on GnRH secretion is at least in part mediated by kisspeptin (KNDy) neurons. Although GnRH neurons express few sex steroid receptors, kisspeptin and KNDy neurons show a high degree of colocalization with estrogen receptors,94 progesterone receptors,153 and androgen receptors.290 In the female monkey, kisspeptin expression is markedly reduced by estrogen or estrogen plus progesterone.91 In addition, sex steroid deficiency is associated with increased kisspeptin expression in the infundibular (arcuate) nucleus—in parallel with circulating gonadotropins—and is reversed by estradiol replacement in females91,94 and by either testosterone or estradiol replacement in males.290 In addition, compared with premenopausal women, the numbers of infundibular prodynorphin-­expressing neurons are decreased in postmenopausal women.291 An autopsy study in men suggests that the numbers of infundibular kisspeptin-­containing cell bodies, fibers, and contacts onto GnRH neurons increase with age, hypothesized to reflect reduced steroid negative feedback.292 In addition to influencing kisspeptin expression, estrogen modulates GnRH neuron responsiveness to kisspeptin in mice.80 A model regarding the influence of KNDy neurons in negative feedback— largely based on data obtained in sheep—is shown in Fig. 1.16. Kisspeptin also plays a key role in the mid-­cycle LH surge in the female rodent. Kiss1 and Kiss1R null mice do not exhibit LH surges,293 and the LH surge can be prevented by a kisspeptin antagonist294 or a monoclonal antibody to kisspeptin.295 Of particular interest, available data in rodents suggest that kisspeptin neurons in the AVPV/RP3V are stimulated by estradiol, whereas those in the arcuate nucleus are inhibited by estradiol.94 In addition, estradiol administration directly into the medial preoptic area (location of the AVPV) induces LH surges, whereas estradiol administration directly into the mediobasal hypothalamus—which also increases estradiol levels in the pituitary—does not.296 These and other findings in rodents have led to a model in which kisspeptin neurons in the arcuate nucleus regulate tonic GnRH release by mediating estrogen negative feedback, whereas

POA

MBH Stress

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Nutrient signals

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GnRH KNDy OC

MB me High E2

E2-Prog inhibition

Fig. 1.16 Model of KNDy signaling to gonadotropin-­releasing hormone (GnRH) neurons, largely based on data obtained in sheep.  KNDy peptides are kisspeptin (green), which stimulates GnRH neurons, neurokinin B (NKB; magenta), and dynorphin (DYN; red). The major influences on GnRH secretion are shown, with putative effects on KNDy peptides denoted by the color of the arrow. For example, estradiol (E2) inhibition may involve reductions of kisspeptin (green arrow), whereas progesterone (Prog) inhibition likely involves an increase in DYN. Arrows with two colors signify that more than one KNDy peptide may mediate a given effect (e.g., in the ewe, stimulation of GnRH secretion by high E2 may involve an increase in both kisspeptin and NKB). The possibility that kisspeptin stimulation of GnRH neurons is mediated by interneurons is shown by the gray cell. MB, Mammillary bodies; MBH, mediobasal hypothalamus; ME, median eminence; OC, optic chiasm; POA, preoptic area. (Modified from Lehman MN, Coolen LM, Goodman RL. Mini review: kisspeptin/ neurokinin B/dynorphin [KNDy] cells of the arcuate nucleus: a central node in the control of gonadotropin-­releasing hormone secretion. Endocrinology. 2010;151:3479–3489.)

kisspeptin neurons in the AVPV/RP3V mediate the positive feedback effects of estrogen (Fig. 1.17). It remains unclear whether a population of kisspeptin neurons homologous to those in the AVPV of rodents plays a similar role in primates. Although LH surge generation requires an intact preoptic area in rodents,297 monkeys retain the ability to produce LH surges after isolation of the mediobasal hypothalamus from the remainder of the brain.178 In addition, LH surges persisted after destruction of the preoptic area, which included the AVPV and suprachiasmatic nuclei in one monkey study,298 but not in another.299 Although human and monkey studies suggest the presence of kisspeptin neurons in the preoptic area,91,92,98,174 it is unclear if these are analogous to those of the AVPV in rodents. A recent study indicated that rostral hypothalamic kisspeptin neuron numbers (which could potentially be homologous to kisspeptin neurons in the rodent RP3V) were lower in postmenopausal women, suggesting positive regulation by estrogens.98 Also of interest, kisspeptin expression increases in a caudal portion of the arcuate nucleus during the preovulatory period in both monkeys and sheep,174,300,301 and some have suggested that kisspeptin neurons in this region could possibly represent a special population important for surge generation. Regardless, it is not certain that a unique population of kisspeptin neurons need be invoked in women, as increased GnRH secretion at mid-­cycle (i.e., a GnRH surge) does not clearly occur in women (described earlier in the chapter). How these intriguing observations in animal models relate to human neurophysiology during mid-­cycle thus remains unclear.

Selective Regulation of Pituitary Follicle-­Stimulating Hormone Secretion Inhibins, activins, and follistatin preferentially influence FSH secretion and contribute to divergent release of LH and FSH During the mid-­to late

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AVPV Kiss1 Neuron



Reproductive Neuroendocrine Adaptations in Settings of Reduced Energy Availability, Stress, and Lactation

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Fig. 1.17 Model of kisspeptin-­mediated feedback regulation of gonadotropin-­releasing hormone (GnRH) and gonadotropin secretion in rodents.  By this model, kisspeptin (Kiss1) neurons in the arcuate nucleus of males and females project to and stimulate GnRH neurons. This population of kisspeptin neurons is inhibited by sex-­appropriate gonadal steroids (i.e., estradiol [E], progesterone [P], and testosterone [T]). Thus, tonic GnRH secretion is primarily regulated by relatively low concentrations of estradiol via kisspeptin neurons in the arcuate nucleus. Females have another population of kisspeptin neurons in the anteroventral periventricular nucleus (AVPV) that also projects to and stimulates GnRH secretion. However, estradiol stimulates kisspeptin neurons in the AVPV—in contrast with estradiol inhibition of kisspeptin neurons in the arcuate nucleus. Thus, although high estradiol concentrations inhibit arcuate kisspeptin neurons in females, they stimulate AVPV kisspeptin neurons, resulting in a GnRH surge. FSH, Follicle-­stimulating hormone; LH, luteinizing hormone. (From Oakley AE, Clifton DK, Steiner RA. Kisspeptin signaling in the brain. Endocr Rev. 2009;30:713–743.)

follicular phase and the luteal phase, both estradiol and inhibins selectively inhibit FSH release from gonadotropes. Inhibins are heterodimer peptide members of the transforming growth factor (TGF)-­β superfamily with two isoforms, inhibin A and inhibin B, which contain identical α-­subunits but different β-­ subunits (βA for inhibin A, βB for inhibin B). Most inhibin is derived from the ovaries: inhibin B is secreted by ovarian granulosa cells, mainly during the early follicular phase in response to FSH stimulation; and inhibin A is primarily produced by the corpus luteum during the luteal phase in response to LH stimulation. The chief function of both inhibins is to inhibit FSH release from pituitary gonadotropes. In men, inhibin B is produced from Sertoli cells and is a key negative feedback regulator of pituitary FSH release, although estradiol also inhibits pituitary FSH release.302 Activin is a dimer peptide with three isoforms: activin A (βAβA), activin B (βBβB), and activin AB (βAβB). The activin β-­subunits and the inhibin β-­subunits are identical. Activin produced in pituitary gonadotropes stimulates production of FSH in a paracrine fashion. Follistatin is a monomer peptide synthesized by the anterior pituitary (including folliculostellate cells); it inhibits pituitary FSH secretion by binding activin, thus rendering it inactive. Interestingly, gonadotrope follistatin production varies in parallel with GnRH pulse frequency, which is one of the mechanisms contributing to the differential effects of GnRH pulse frequency on LH and FSH release.57,303 In contrast to inhibins, which act primarily via endocrine signaling, activin and follistatin produced in the pituitary influence FSH secretion via autocrine-­paracrine signaling.

• Reproductive function is impaired in the setting of decreased energy availability and/or stress; this primarily reflects central inhibition of GnRH pulse frequency and reduced gonadotropin secretion. • Lactation is associated with suppressed pulsatile GnRH secretion and low gonadotropin concentrations; this relates to the high energy demands of lactation (reduced energy availability), hyperprolactinemia, and other (e.g., neural) mechanisms.

Interface Between Reproductive Neuroendocrine Function and Energy Availability Organisms require metabolic energy to support a number of processes, including maintenance of cellular function, muscle contraction (e.g., cardiac function, locomotion), thermogenesis, and growth. Low energy availability may result from short-­or long-­ term reductions in calorie intake (e.g., famine, anorexia nervosa), insufficient calorie intake for metabolic demands (e.g., in the setting of significant exercise loads or hypermetabolic states), or reduced ability to use energy sources (e.g., as may occur in severe diabetes). In such situations, energy use has opportunity costs; energy used for one process is no longer available for another. Energy-­requiring processes are thus prioritized to favor those that are life sustaining. Reproduction in women, pregnancy and lactation in particular, is metabolically demanding. For example, pregnancy requires an estimated additional 80,000 kilocalories.304 Because reproduction is not imperative for individual survival, it is metabolically gated; reductions in energy availability can suppress reproductive function (nutritional infertility). This is biologically advantageous for the individual and, ultimately, for the species. As such, it can be seen as an appropriate adaptive response. This process is believed to be at the center of functional hypothalamic amenorrhea, a reversible condition of suppressed hypothalamic-­pituitary function occurring in the absence of anatomic abnormalities and often accompanied by reduced body weight, disordered eating (e.g., restrictive eating patterns), excessive exercise, and/or psychological stress. The functional relationships between metabolic status and reproductive function are mediated by neural systems located in the hypothalamus, and functional hypothalamic amenorrhea is characterized by impaired GnRH and gonadotropin secretion. Although a majority of women with functional hypothalamic amenorrhea demonstrate low LH pulse frequency,305,306 such patients may demonstrate variable LH pulse patterns as a group. Such patterns include absent pulses, low frequency and amplitude, low frequency only, low amplitude only, and (apparently) normal frequency and amplitude. Patterns can change across time in the same woman.8,305–307 In these patients, GnRH and gonadotropin secretion is inadequate for normal follicular development, estrogen production, and mid-­cycle gonadotropin surges, but cyclic ovulation and fertility can be restored with pulsatile administration of exogenous GnRH.8 Some investigators have proposed that reduced reproductive function in functional hypothalamic amenorrhea primarily reflects insufficient body fat stores—the critical fatness hypothesis.308 However, a substantial body of data suggests that reduced energy availability is the primary cause of reduced reproductive function in these settings. For example, body fat does not reliably distinguish amenorrheic from eumenorrheic athletes.309 In addition, calorie restrictions can result in amenorrhea before substantial weight loss, and amenorrhea can persist after weight restoration in those with a history of eating disorders.310 Likewise,

CHAPTER 1  Neuroendocrinology of Reproduction 45

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Fig. 1.18 Influence of energy availability on luteinizing hormone (LH) pulsatility in healthy women.  (A) Representative 24-­hour LH time series in three women under different conditions of energy availability. These studies were designed to assess the effects of exercise and reduced energy availability in habitually sedentary women with regular menstrual cycles. In this study, exercise energy expenditure was substantial for all participants at approximately 840 kcal/day, and energy availability was altered via variable calorie intake. When energy availability is defined as dietary energy intake minus exercise energy expenditure—an estimate of the energy available for non-­exercise-­related functions—and normalized to fat-­free mass (thus expressed as kcal/kg lean body mass [LBM] per day), 45 kcal/kg LBM per day approximates balanced energy availability, and LH profiles under this condition are shown on top. Conditions of restricted energy availability (i.e., 10, 20, and 30 kcal/kg LBM per day) are shown along the bottom. Significant LH pulses are denoted by asterisks; arrows denote the timing of meals; and black bars denote lights out periods. (B) Association of energy availability and LH pulse characteristics. Energy availability is shown on the x-­axis; on the y-­axis, LH pulse amplitude (solid circles) and LH pulse frequency (open circles) are expressed as changes relative to values observed at 45 kcal/kg LBM per day. (Note that changes of LH pulse amplitude are divided by three.) Although energy availability reductions to 30 kcal/kg LBM per day did not alter LH pulse characteristics, reductions below 30 kcal/kg LBM per day were associated with progressive reductions of LH pulse frequency and corresponding increases of LH pulse amplitude. (Modified from Loucks AB, Thuma JR. Luteinizing hormone pulsatility is disrupted at a threshold of energy availability in regularly menstruating women. J Clin Endocrinol Metab. 2003;88:297–311.)

findings consistent with functional hypothalamic amenorrhea can be observed shortly after bariatric surgery for severe obesity in the setting of negative energy balance but while still obese (e.g., body mass index approximately 35 kg/m2).311 Moreover, altered LH pulsatility is observed very quickly, within 5 days of controlled reductions of energy availability,312 although experiments in women and monkeys suggest that reproductive function may not be impaired until energy availability is reduced by more than 30% (Fig. 1.18).312,313 Overall, these findings suggest that altered reproductive function in this setting reflects reduced energy availability rather than reduced body fat stores per se. It is important to note that energy balance (and thus body weight)

can be maintained in the face of calorie restriction by reducing metabolic rate314 and by suspending “noncritical” but energy-­ requiring functions such as reproduction. Although some have posited a specific influence of exercise, calorie supplementation to maintain adequate energy availability appears to prevent alterations of LH secretion despite significant daily exercise loads.315 Similarly, whereas amenorrhea can be induced in monkeys by gradually increasing daily exercise in the setting of constant food intake,316 providing supplemental calories reverses amenorrhea despite continued exercise.317 The neurobiologic mechanisms underlying functional hypothalamic amenorrhea, and mechanisms underlying the

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PART I  The Fundamentals of Reproduction

influence of nutritional status on the reproductive system in general, remain poorly understood. Chronic energy deprivation is associated with myriad neuroendocrine adaptations and hormonal changes, including reductions of circulating leptin, insulin, insulin-­like growth hormone-­1 (IGF-­1), and thyroid hormone concentrations, as well as increases of growth hormone and ghrelin levels and activation of the hypothalamic-­ pituitary-­adrenal (HPA) axis.318 A number of these alterations can influence GnRH and gonadotropin secretion and may mediate the influence of low energy availability on reproductive function. Much interest has focused on the permissive role of leptin, a hormone derived from adipose tissue that functions to signal metabolic status to central systems, influencing feeding, energy expenditure, and reproduction. Humans and mice lacking leptin or the leptin receptor (LepR) (e.g., ob/ob and db/db mice, respectively) have pubertal failure and infertility, and the reproductive manifestations of leptin deficiency can be reversed with leptin administration.319,320 Low leptin levels have also been observed in women with functional hypothalamic amenorrhea,321 and a specific role for leptin is suggested by studies in which recombinant human leptin administration improved LH pulse secretion, estradiol concentrations, and menstrual cyclicity, at least in a proportion of women with functional hypothalamic amenorrhea.322–324 Although serum leptin concentrations generally correlate with fat mass, leptin levels can change rapidly and are suppressed with maneuvers known to suppress LH secretion, such as marked short-­term energy restriction.325 A number of neuropeptides have been implicated in the influence of energy availability on reproductive function, including kisspeptin, NPY, galanin-­ like peptide (GALP), β-­ endorphin, CRH, ghrelin, and peptide YY. Although metabolic signals could potentially act directly on GnRH neurons, many studies suggest that afferent neural circuits are involved. For example, studies in mice suggest that metabolic signals can be relayed to the GnRH pulse generator from different areas in the brain, such as the ventral premammillary nucleus (e.g., leptin effects)326 and the area postrema (e.g., in the absence of usable glucose).327 A growing body of evidence suggests that the influence of energy availability on reproductive neuroendocrine function is at least partly mediated by kisspeptin neurons.328,329 For example, leptin-­deficient (ob/ob) mice demonstrate reduced hypothalamic Kiss1 expression, which is partially reversed with leptin administration.330 Some studies suggest that increased opioid tone contributes to slow GnRH pulses in functional hypothalamic amenorrhea.306,331–333 Although successful reproduction is not metabolically costly for males, male reproductive function can also be disrupted by metabolic stress. For example, healthy young men participating in US Army Ranger training (which involves multiple stressors, including intermittent extreme calorie restriction and weight loss [10–12 kg on average]) can experience reductions in LH secretion and suppression of testosterone concentrations to near castrate levels.334 Increased calorie intake allowed prompt recovery of testosterone in this study, even without altering other associated stressors (e.g., exercise, sleep deprivation). In addition, anorexia nervosa in adolescent boys and men can be associated with marked hypogonadotropic hypogonadism.335 Although men appear to be less likely than women to experience disrupted reproductive function in the setting of metabolic stress, a form of reversible, functional hypogonadotropic hypogonadism related to excessive exercise and/or weight loss has been described in men, which is in many ways analogous to functional hypothalamic amenorrhea in women.336 Also of interest, exogenous leptin administration prevented the fall of LH release and testosterone concentrations associated with short-­term fasting in men.337 Energy-­sensitive reproductive function in males would also be expected to delay reproduction during times of reduced energy availability; these

regulatory pathways may have developed in males because of advantages imparted to other members of the species (e.g., mates and offspring).

Impact of Stress on Reproductive Neuroendocrine Function Functional hypothalamic amenorrhea in the setting of reduced energy availability represents a particular form of stress-­related reproductive suppression. The term stressor refers to a real or potential threat to homeostasis, such as injury, illness, temperature extremes, reduced energy availability, predator proximity, and situations that provoke psychological distress. The nature of the stress response depends on the precise nature of the stressor but typically involves both neural and neuroendocrine responses. A group of neurons in the hypothalamic paraventricular nucleus project to the median eminence, where they secrete CRH into the hypophyseal portal system. CRH (and, to some degree, cosecreted AVP) stimulates corticotrope cells in the anterior pituitary to release ACTH, which in turn stimulates adrenal glucocorticoid (cortisol) production. A subset of paraventricular neurons is also involved with the regulation of the autonomic sympathetic nervous system, which includes neural pathways linked to the brain stem, spinal cord, and adrenal medulla (e.g., the sympathoadrenal axis). Other components of the stress response include central arousal systems and the locus ceruleus, a nucleus in the brainstem involved with emotional and cognitive responses to stress. Thus, stressors trigger integrated neural, endocrine, and behavioral responses that promote short-­term maintenance of homeostasis and survival. For example, activation of the sympathoadrenal axis leads to increased epinephrine secretion—an important component of the fight-­ or-­ flight response—whereas activation of the HPA axis with increased cortisol secretion enhances energy mobilization. Chronic stress and marked acute stress can inhibit reproductive function—an appropriate adaptive response when homeostasis is threatened. For example, critical illness is associated with reversible hypogonadotropic hypogonadism.338,339 Mechanisms underlying the suppression of reproductive function during stress are highly complex; although suppression of GnRH secretion is a major component, direct pituitary and gonadal effects may also occur. Notably, the specific effects of stress on various aspects of reproductive function appear to depend on a number of factors including species, sex, hormonal milieu (e.g., gonad intact vs. castrate), and the specific type of stress experienced. A number of mediators have been implicated in stress-­ related inhibition of GnRH secretion, including CRH (which activates the HPA axis but also appears to have central effects), CRH-­ like peptides called urocortins, AVP, ACTH, EOPs (e.g., β-­endorphin), and cortisol, in addition to noradrenergic, GABAergic, and serotoninergic neural pathways. For example, intracerebroventricular injection of CRH reduces multiple unit electrical activity in the mediobasal hypothalamus in monkeys,340 and CRH antagonists can prevent some forms of stress-­related LH suppression.341 Naloxone can block CRH-­related LH suppression in monkeys,342 suggesting the involvement of EOPs in this process. Notably, some data suggest that stress plays a role in functional hypothalamic amenorrhea. For example, amenorrheic athletes and women with anorexia nervosa demonstrate evidence of HPA axis activation (e.g., elevated cortisol concentrations).318,343 In addition, functional hypothalamic amenorrhea in women may be associated with evidence of higher psychological stress, including perfectionism, a history of unfavorable childhood experiences, and difficulty coping with stressors.344,345 Moreover, behavioral therapy346 or hypnotherapy347 may improve reproductive function in some women with functional hypothalamic amenorrhea. Studies in female monkeys provide corroborating evidence: in one

CHAPTER 1  Neuroendocrinology of Reproduction

study, very few monkeys demonstrated altered reproductive function when exposed to either (1) psychosocial stress (relocation to new housing setting with unfamiliar monkeys) or (2) mild dietary restriction plus daily exercise, but the combination was associated with altered cycle length or anovulation in a majority.348 For unclear reasons, the degree to which stressors (e.g., reduced energy availability) interrupt reproductive function is variable among individual women (i.e., hypothalamus robustus vs. hypothalamus fragilis; stress sensitive vs. stress resilient). As suggested previously, it is likely that a number of factors (e.g., reduced energy availability, stress) can interact to impact GnRH secretion. In addition, two studies have suggested that variants in genes associated with hypogonadotropic hypogonadism—including ANOS1, PROKR2, and the GnRH receptor (GNRHR) genes, among others—are more likely to be identified in women with functional hypothalamic amenorrhea compared with normally cycling women.349,350 Thus, it seems likely that underlying genetic (and epigenetic) architecture plays an important role in reproductive susceptibility to reduced energy availability and stress.

Lactation and Reproductive Neuroendocrine Function High prolactin concentrations during pregnancy and suckling in the postpartum period stimulate milk production, which, for much of human history, was effectively the only source of nutrition for infants. Suckling also leads to posterior pituitary release of oxytocin, which stimulates contraction of myoepithelial cells within mammary gland acini, causing milk ejection. Lactation is associated with amenorrhea and subfertility. The likelihood of pregnancy during the first 6 months postpartum is low (less than 2%) in fully breastfeeding, amenorrheic women,351 and some lactating women may remain amenorrheic for years. Because a short interval between births can place infant well-­ being at risk, lactational amenorrhea has ostensibly been an important adaptation enhancing infant survival in many cultures both past and present.352 During pregnancy, high placental sex steroid (estradiol, progesterone) and prolactin concentrations markedly suppress GnRH and gonadotropin secretion and prevent follicular development. In the absence of lactation, cyclic hypothalamic-­ pituitary-­ovarian activity typically resumes within 8 weeks after parturition. However, in the setting of lactation, pulsatile GnRH remains suppressed (e.g., low-­frequency pulses) with consequent impairment of LH secretion and estradiol production.353 The reduction in GnRH secretion during lactation is suggested by a marked reduction in multiple unit electrical activity in the mediobasal hypothalamus in nursing monkeys354 and by the ability of pulsatile exogenous GnRH to restore ovarian function in amenorrheic lactating women.355 Mechanisms underlying lactational amenorrhea are not completely understood. Lactation is associated with a very high metabolic cost; daily production of 750 to 1000 mL of human milk requires approximately 500 to 600 kilocalories a day,356 some of which is obtained from fat stores and increased food intake. Nonetheless, the high energy requirements of lactation, which are approximately twice that of pregnancy, may induce some or all of the aforementioned mechanisms that inhibit pulsatile GnRH secretion in the setting of reduced energy availability. Animal (chiefly rodent) studies reveal that lactation is associated with activation of orexigenic neural systems (e.g., NPY) and inhibition of anorexigenic neural systems within the hypothalamus (changes that may partly relate to alterations of peripheral metabolic cues, such as leptin and insulin.357 Such alterations increase food intake and may suppress GnRH neuronal activity, either through direct effects on the GnRH neuronal network or modification of key afferent systems such as kisspeptin. As an example of the former, NPY neuronal activation during lactation may directly inhibit

21

GnRH neuronal activity in rats.358 As an example of the latter, expression of both kisspeptin and NKB in the arcuate nucleus is reduced in lactating rodents.161,359 Hyperprolactinemia suppresses GnRH pulsatility, at least in part via increased hypothalamic opioids,360 and suppression of GnRH secretion during periods of lactation may partly reflect high prolactin concentrations. Data in rodent models suggest that prolactin suppression of GnRH secretion is at least partly mediated by kisspeptin: kisspeptin neurons appear to express prolactin receptors361; hyperprolactinemia has been associated with reduced hypothalamic Kiss1 expression362,363; prolactin does not impair LH release in mice after selective prolactin receptor-­ knockout of arcuate nucleus kisspeptin neurons364; and kisspeptin administration reverses hyperprolactinemia-­ mediated hypogonadotropic anovulation.362 Moreover, in women with hyperprolactinemia, exogenous kisspeptin partly restores gonadotropin secretion and estradiol concentrations.110 However, prolactin levels gradually decrease to normal despite continued breastfeeding; thus, the activity of the hypothalamic-­pituitary-­ovarian axis may not correlate well with circulating prolactin concentrations. The intensity (frequency and duration) of suckling appears to be an important determinant of contraceptive effectiveness in women, and the suckling stimulus may inhibit the hypothalamic-­pituitary-­ ovarian axis through additional (e.g., neural) mechanisms.

Miscellaneous Physiologic Influences on Gonadotropin-­Releasing Hormone Secretion Circadian Changes Diurnal rhythms (i.e., those that cycle once a day) are frequently observed in endocrinology, including the reproductive system. Diurnal rhythms are called circadian rhythms if they are internally (endogenously) driven rhythms, although such rhythms are usually entrained (synchronized) to environmental cues (e.g., light-­dark cycle). Circadian rhythms are dictated by a “master clock” located in the suprachiasmatic nucleus of the hypothalamus and are derived from complex intracellular interactions involving the so-­called clock genes, which participate in feedback interactions that generate recurring cyclic activity.365 Such clocks in the suprachiasmatic nucleus can be synchronized by light-­dark signals received from the retina and transferred to the suprachiasmatic nucleus via the optic nerve (retinohypothalamic tract). Depending on such factors as species and sex, both basal gonadotropin secretion and LH surges may exhibit diurnal rhythms. As a prominent example, LH surges in female rats are specifically confined to the late afternoon, shortly before rats become active (i.e., when copulation is most likely).366 It is believed that this reflects a daily stimulus generated by the suprachiasmatic nucleus but relayed to the GnRH neuronal network only in the presence of preovulatory estradiol concentrations. Thus ovulation in rats is optimally timed to coincide with sexual opportunity and receptivity. In contrast to rodents, LH surges do not appear to be constrained to a specific time of day in monkeys; for example, LH surges can be advanced by 12 to 18 hours with supraphysiologic estradiol administration.244 Some studies in women suggest that LH surges tend to be initiated in the morning.367,368 For example, in one study of 19 ovulatory women, LH surges were initiated in the early morning hours (approximately 4:00 to 8:00 a.m.)367; in one study of 155 spontaneous cycles, the estimated time of LH surge initiation was between midnight and 8:00 a.m. in 85% of cycles.368 In contrast to these findings, a detailed study of mid-­ cycle gonadotropin surges in women suggested that surge initiation is not constrained to a certain time of day.264 In addition, the potential relevance of a specific daily timing of ovulation in women is uncertain; ovulation typically occurs some 36 hours

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PART I  The Fundamentals of Reproduction

after the LH surge, and the likelihood of conception when sexual intercourse occurs on the day before ovulation (approximately 40%) is similar to conception rates when intercourse occurs on the day of ovulation.369 Humans demonstrate diurnal changes of gonadotropins and sex steroids. For example, serum LH concentrations in cycling women tend to peak in the afternoon and reach a nadir at night, particularly in the early follicular phase.219,370,371 Similar findings pertain to serum FSH concentrations.372 However, such findings do not confirm a true circadian signal, and diurnal changes could reflect environmental cues and the influence of sleep. As a prominent example, LH concentrations decline during sleep in the early follicular phase, regardless of whether sleep occurs during nighttime or daytime hours.373 Although one study of cycling women studied under constant-­routine conditions suggested an underlying circadian rhythm for circulating LH and FSH levels during the follicular phase (and FSH during the luteal phase), 374 other studies in women who were carefully assessed during the early follicular phase375 or after menopause376 suggested that LH and FSH secretory parameters (including LH pulse frequency and amplitude) do not exhibit circadian changes after controlling for sleep status, body position, light exposure, activity level, and nutritional cues.

Sleep Although precise mechanisms are unclear, sleep can have a major influence on pulsatile LH secretion (and GnRH by inference). For example, the nocturnal amplification of LH pulsatility during puberty is specifically related to sleep; it generally begins within an hour of sleep onset and follows sleep reversal.377,378 Although the data relating LH pulses to sleep stage are incomplete, early studies suggested that sleep-­related pulses during puberty occur primarily during non-­rapid eye movement (REM) sleep.377,378 Further refining this concept, more recent studies suggest a strong relationship between slow wave sleep and LH pulse initiation during puberty.379,380 Sleep also influences LH pulse secretion in adult women, primarily in the form of sleep-­ related slowing of LH pulse frequency, which is most prominent during the early follicular phase,217,219,371,381 but also occurs in the late follicular phase.312,382 Nocturnal slowing of LH pulse frequency in women during the early follicular phase is specifically related to sleep: it accompanies daytime sleep, and it is absent during nighttime wakefulness.373,375,383 During the follicular phase, LH pulses are uncommon during REM and slow wave sleep and more common following brief awakenings.383,384 Such slowing may be mediated by hypothalamic opioids because naloxone appeared to prevent the sleep-­associated decrease in LH pulse frequency.370 Interestingly, sleep appears to interact with other determinants of pulsatile GnRH secretion. For example, studies in peripubertal girls suggest that progesterone acutely suppresses LH pulse frequency during waking hours but not during nighttime (sleeping) hours.385,386 Similarly, studies performed during the late follicular phase in normal women suggested that dietary calorie restriction preferentially reduces daytime LH pulse frequency.315,382 These findings suggest differential control of GnRH pulse frequency depending on sleep status in human females. Taken together, the aforementioned data imply that sleep influences LH pulse frequency and that the effect of sleep can be modulated by such factors as developmental stage and sex steroid milieu. The physiologic relevance of sleep-­associated changes of GnRH secretion remains unclear, but such changes have been postulated to contribute to normal gonadotropin production across puberty203 and to the prominence of FSH secretion during the early follicular phase in postpubertal women.383

Pheromones Pheromones are chemicals transmitted through the air that can influence reproductive function and sexual behavior in other individuals within many mammalian species. For example, the presence of a novel and sexually mature male mouse can synchronize estrous cycles among female mice; this so-­called Whitten effect is presumably mediated by pheromones.387 Similarly, pheromones produced by male sheep and goats can induce out-­of-­season ovulation in females; this is called the male effect.388 The role of pheromones in humans remains unclear. Menstrual synchrony—sometimes called the McClintock effect—is a putative phenomenon in which menstrual cycles of women living in close proximity become synchronized389; this has been cited as physiologic evidence of pheromone functionality in humans. Axillary (armpit) compounds obtained from women during the late follicular phase have been reported to advance ovulatory timing in recipient women.390 However, supportive research has been criticized on methodologic grounds, and the existence of this phenomenon remains controversial.391 In addition, although the vomeronasal organ—believed to be responsible for pheromone detection in animals—develops in utero in humans, it subsequently regresses and is largely believed to be nonfunctional in adults.392 TOP REFERENCES

Boehm U, Bouloux P-­M, Dattani MT, et al. Expert consensus document: European Consensus Statement on congenital hypogonadotropic hypogonadism—Pathogenesis, diagnosis and treatment. Nat Rev Endocrinol. 2015;11:547–564. Clarke SA, Dhillo WS. Kisspeptin across the human lifespan: evidence from animal studies and beyond. J Endocrinol. 2016;229:R83–R98. Garcia JP, Keen KL, Seminara SB, Terasawa E. Role of kisspeptin and NKB in puberty in nonhuman primates: sex differences. Semin Reprod Med. 2019;37(2):47–55. Herbison AE. Control of puberty onset and fertility by gonadotropin-­ releasing hormone neurons. Nat Rev Endocrinol. 2016;12:452–466. Herbison AE. The gonadotropin-­ releasing hormone pulse generator. Endocrinology. 2018;159:3723–3736. Hunjan T, Abbara A. Clinical translational studies of kisspeptin and neurokinin B. Semin Reprod Med. 2019;37:119–124. Lehman MN, He W, Coolen LM, Levine JE, Goodman RL. Does the KNDy model for the control of gonadotropin-­releasing hormone pulses apply to monkeys and humans? Semin Reprod Med. 2019;37:71–83. McCosh RB, Breen KM, Kauffman AS. Neural and endocrine mechanisms underlying stress-­induced suppression of pulsatile LH secretion. Mol Cell Endocrinol. 2019;498:110579. McNeilly AS. Neuroendocrine changes and fertility in breast-­feeding women. Prog Brain Res. 2001;133:207–214. Moore AM, Coolen LM, Porter DT, Goodman RL, Lehman MN. KNDy cells revisited. Endocrinology. 2018;159:3219–3234. Navarro VM, Kaiser UB. Metabolic influences on neuroendocrine regulation of reproduction. Curr Opin Endocrinol Diabetes Obes. 2013;20:335–341. Plant TM. A comparison of the neuroendocrine mechanisms underlying the initiation of the preovulatory LH surge in the human, old world monkey and rodent. Front Neuroendocrinol. 2012;33:160–168. Plant TM. The neurobiological mechanism underlying hypothalamic GnRH pulse generation: the role of kisspeptin neurons in the arcuate nucleus. F1000Res. 2019;8:982. Skorupskaite K, George JT, Anderson RA. The kisspeptin-­GnRH pathway in human reproductive health and disease. Hum Reprod Update. 2014;20:485–500. Terasawa E. Mechanism of pulsatile GnRH release in primates: Unresolved questions. Mol Cell Endocrinol. 2019;498:110578.

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REFERENCES

1. Herde MK, Iremonger KJ, Constantin S, Herbison AE. GnRH neurons elaborate a long-­range projection with shared axonal and dendritic functions. J Neurosci. 2013;33(31):12689–12697. 2. Ojeda SR, Lomniczi A, Sandau U. Contribution of glial-­neuronal interactions to the neuroendocrine control of female puberty. Eur J Neurosci. 2010;32:2003–2010. 3. Prevot V. Glial-­ neuronal-­ endothelial interactions are involved in the control of GnRH secretion. J Neuroendocrinol. 2002;14: 247–255. 4. Schally AV, Arimura A, Kastin AJ, et al. Gonadotropin-­releasing hormone: one polypeptide regulates secretion of luteinizing and follicle-­stimulating hormones. Science. 1971;173:1036. 1036. 5. Yu B, Ruman J, Christman G. The role of peripheral gonadotropin-­ releasing hormone receptors in female reproduction. Fertil Steril. 2011;95:465–473. 6. Mason AJ, Pitts SL, Nikolics K, et al. The hypogonadal mouse: reproductive functions restored by gene therapy. Science. 1986;234:1372–1373. 7. Gibson MJ, Krieger DT, Charlton HM, Zimmerman EA, Silverman AJ, Perlow MJ. Mating and pregnancy can occur in genetically hypogonadal mice with preoptic area brain grafts. Science. 1984;225:949–951. 8. Santoro N, Filicori M, Crowley WF Jr. Hypogonadotropic disorders in men and women: diagnosis and therapy with pulsatile gonadotropin-­releasing hormone. Endocr Rev. 1986;7:11–23. 9. Millar RP, Lu ZL, Pawson AJ, Flanagan CA, Morgan K, Maudsley SR. Gonadotropin-­ releasing hormone receptors. Endocr Rev. 2004;25:235–275. 10. Cheng CK, Leung PC. Molecular biology of gonadotropin-­ releasing hormone (GnRH)-­ I, GnRH-­ II, and their receptors in humans. Endocr Rev. 2005;26:283–306. 11. Rance NE, Young 3rd WS, McMullen NT. Topography of neurons expressing luteinizing hormone-­releasing hormone gene transcripts in the human hypothalamus and basal forebrain. J Comp Neurol. 1994;339:573–586. 12. Stopa EG, Koh ET, Svendsen CN, Rogers WT, Schwaber JS, King JC. Computer-­ assisted mapping of immunoreactive mammalian gonadotropin-­releasing hormone in adult human basal forebrain and amygdala. Endocrinology. 1991;128:3199–3207. 13. Plant TM, Krey LC, Moossy J, McCormack JT, Hess DL, Knobil E. The arcuate nucleus and the control of gonadotropin and prolactin secretion in the female rhesus monkey (Macaca mulatta). Endocrinology. 1978;102:52–62. 14. Campbell RE, Gaidamaka G, Han SK, Herbison AE. Dendro-­ dendritic bundling and shared synapses between gonadotropin-­ releasing hormone neurons. Proc Natl Acad Sci USA. 2009;106:10835–10840. 15. Moore AM, Prescott M, Czieselsky K, et al. Synaptic innervation of the GnRH neuron distal dendron in female mice. Endocrinology. 2018;159:3200–3208. 16. Fueshko S, Wray S. LHRH cells migrate on peripherin fibers in embryonic olfactory explant cultures: an in vitro model for neurophilic neuronal migration. Dev Biol. 1994;166:331–348. 17. Bless EP, Walker HJ, Yu KW, et al. Live view of gonadotropin-­ releasing hormone containing neuron migration. Endocrinology. 2005;146:463–468. 18. Schwanzel-­Fukuda M, Pfaff DW. Origin of luteinizing hormone-­ releasing hormone neurons. Nature. 1989;338:161–164. 19. Wray S, Nieburgs A, Elkabes S. Spatiotemporal cell expression of luteinizing hormone-­releasing hormone in the prenatal mouse: evidence for an embryonic origin in the olfactory placode. Brain Res Dev Brain Res. 1989;46:309–318. 20. Wray S, Grant P, Gainer H. Evidence that cells expressing luteinizing hormone-­releasing hormone mRNA in the mouse are derived from progenitor cells in the olfactory placode. Proc Natl Acad Sci U S A. 1989;86:8132–8136. 21. Ronnekleiv OK, Resko JA. Ontogeny of gonadotropin-­releasing hormone-­containing neurons in early fetal development of rhesus macaques. Endocrinology. 1990;126:498–511. 22. Kim KH, Patel L, Tobet SA, King JC, Rubin BS, Stopa EG. Gonadotropin-­ releasing hormone immunoreactivity in the adult and fetal human olfactory system. Brain Res. 1999;826:220–229. 23. Livne I, Gibson MJ, Silverman AJ. Biochemical differentiation and intercellular interactions of migratory gonadotropinmone (GnRH) cells in the mouse. . 1993;159:643–656.

24. Yoshida K, Tobet SA, Crandall JE, Jimenez TP, Schwarting GA. The migration of luteinizing hormone-­releasing hormone neurons in the developing rat is associated with a transient, caudal projection of the vomeronasal nerve. J Neurosci. 1995;15:7769–7777. 25. Boehm U, Bouloux PM, Dattani MT, et al. Expert consensus document: European Consensus Statement on congenital hypogonadotropic hypogonadism—Pathogenesis, diagnosis and treatment. Nat Rev Endocrinol. 2015;11:547–564. 26. Schwanzel-­Fukuda M, Bick D, Pfaff DW. Luteinizing hormone-­ releasing hormone (LHRH)-­expressing cells do not migrate normally in an inherited hypogonadal (Kallmann) syndrome. Brain Res Mol Brain Res. 1989;6:311–326. 27. Quinton R, Hasan W, Grant W, et al. Gonadotropin-­releasing hormone immunoreactivity in the nasal epithelia of adults with Kallmann’s syndrome and isolated hypogonadotropic hypogonadism and in the early midtrimester human fetus. J Clin Endocrinol Metab. 1997;82:309–314. 28. Dodé C, Teixeira L, Levilliers J, et al. Kallmann syndrome: mutations in the genes encoding prokineticin-­2 and prokineticin receptor-­2. PLoS Genet. 2006;2:e175. 29. Dodé C, Levilliers J, Dupont JM, et al. Loss-­of-­function mutations in FGFR1 cause autosomal dominant Kallmann syndrome. Nat Genet. 2003;33:463–465. 30. Falardeau J, Chung WC, Beenken A, et al. Decreased FGF8 signaling causes deficiency of gonadotropin-­releasing hormone in humans and mice. J Clin Investig. 2008;118:2822–2831. 31. Miura K, Acierno JS Jr, Seminara SB. Characterization of the human nasal embryonic LHRH factor gene, NELF, and a mutation screening among 65 patients with idiopathic hypogonadotropic hypogonadism (IHH). J Hum Genet. 2004;49:265–268. 32. Kim HG, Kurth I, Lan F, et al. Mutations in CHD7, encoding a chromatin-­ remodeling protein, cause idiopathic hypogonadotropic hypogonadism and Kallmann syndrome. Am J Hum Genet. 2008;83:511–519. 33. Young J, Metay C, Bouligand J, et al. SEMA3A deletion in a family with Kallmann syndrome validates the role of semaphorin 3A in human puberty and olfactory system development. Hum Reprod. 2012;27:1460–1465. 34. Pingault V, Bodereau V, Baral V, et al. Loss-­of-­function mutations in SOX10 cause Kallmann syndrome with deafness. Am J Hum Genet. 2013;92:707–724. 35. Ng KL, Li JD, Cheng MY, Leslie FM, Lee AG, Zhou QY. Dependence of olfactory bulb neurogenesis on prokineticin 2 signaling. Science. 2005;308:1923–1927. 36. Sykiotis GP, Plummer L, Hughes VA, et al. Oligogenic basis of isolated gonadotropin-­releasing hormone deficiency. Proc Natl Acad Sci USA. 2010;107:15140–15144. 37. Raivio T, Falardeau J, Dwyer A, et al. Reversal of idiopathic hypogonadotropic hypogonadism. N Engl J Med. 2007;357:863–873. 38. Sidhoum VF, Chan YM, Lippincott MF, et al. Reversal and relapse of hypogonadotropic hypogonadism: resilience and fragility of the reproductive neuroendocrine system. J Clin Endocrinol Metab. 2014;99:861–870. 39. Levine JE, Duffy MT. Simultaneous measurement of luteinizing hormone (LH)-­ releasing hormone, LH, and follicle-­ stimulating hormone release in intact and short-­term castrate rats. Endocrinology. 1988;122:2211–2221. 40. Levine JE, Pau KY, Ramirez VD, Jackson GL. Simultaneous measurement of luteinizing hormone-­releasing hormone and luteinizing hormone release in unanesthetized, ovariectomized sheep. Endocrinology. 1982;111:1449–1455. 41. Carmel PW, Araki S, Ferin M. Pituitary stalk portal blood collection in rhesus monkeys: evidence for pulsatile release of gonadotropin-­ releasing hormone (GnRH). Endocrinology. 1976;99:243–248. 42. Wildt L, Hausler A, Marshall G, et al. Frequency and amplitude of gonadotropin-­releasing hormone stimulation and gonadotropin secretion in the rhesus monkey. Endocrinology. 1981;109:376–385. 43. Belchetz PE, Plant TM, Nakai Y, Keogh EJ, Knobil E. Hypophysial responses to continuous and intermittent delivery of hypothalamic gonadotropin-­releasing hormone. Science. 1978;202:631–633. 44. Ortmann O, Weiss JM, Diedrich K. Gonadotrophin-­releasing hormone (GnRH) and GnRH agonists: mechanisms of action. Reprod Biomed Online. 2002;5(suppl 1):1–7. 45. Savoy-­Moore RT, Schwartz NB, Duncan JA, Marshall JC. Pituitary releasing hormone receptors during the rat estrous cycle.

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46. Stanton PG, Burgon PG, Hearn MT, Robertson DM. Structural and functional characterisation of hFSH and hLH isoforms. Mol Cell Endocrinol. 1996;125:133–141. 47. Marshall JC, Case GD, Valk TW, Corley KP, Sauder SE, Kelch RP. Selective inhibition of follicle-­stimulating hormone secretion by estradiol. Mechanism for modulation of gonadotropin responses to low dose pulses of gonadotropin-­ releasing hormone. J Clin Investig. 1983;71:248–257. 48. Bilezikjian LM, Blount AL, Leal AM, Donaldson CJ, Fischer WH, Vale WW. Autocrine/paracrine regulation of pituitary function by activin, inhibin and follistatin. Mol Cell Endocrinol. 2004;225:29–36. 49. Clarke IJ, Cummins JT, Findlay JK, Burman KJ, Doughton BW. Effects on plasma luteinizing hormone and follicle-­ stimulating hormone of varying the frequency and amplitude of gonadotropin-­ releasing hormone pulses in ovariectomized ewes with hypothalamo-­ pituitary disconnection. Neuroendocrinology. 1984;39:214–221. 50. Gross KM, Matsumoto AM, Bremner WJ. Differential control of luteinizing hormone and follicle-­stimulating hormone secretion by luteinizing hormone-­releasing hormone pulse frequency in man. J Clin Endocrinol Metab. 1987;64:675–680. 51. Spratt DI, Finkelstein JS, Butler JP, Badger TM, Crowley WF Jr. Effects of increasing the frequency of low doses of gonadotropin-­ releasing hormone (GnRH) on gonadotropin secretion in GnRH-­ deficient men. J Clin Endocrinol Metab. 1987;64:1179–1186. 52. Dalkin AC, Haisenleder DJ, Ortolano GA, Ellis TR, Marshall JC. The frequency of gonadotropin-­releasing-­hormone stimulation differentially regulates gonadotropin subunit messenger ribonucleic acid expression. Endocrinology. 1989;125:917–924. 53. Burger LL, Haisenleder DJ, Dalkin AC, Marshall JC. Regulation of gonadotropin subunit gene transcription. J Mol Endocrinol. 2004;33:559–584. 54. Thackray VG, Mellon PL, Coss D. Hormones in synergy: regulation of the pituitary gonadotropin genes. Mol Cell Endocrinol. 2010;314:192–203. 55. Stamatiades GA, Kaiser UB. Gonadotropin regulation by pulsatile GnRH: signaling and gene expression. Mol Cell Endocrinol. 2018;463:131–141. 56. Bedecarrats GY, Kaiser UB. Differential regulation of gonadotropin subunit gene promoter activity by pulsatile gonadotropin-­releasing hormone (GnRH) in perifused L beta T2 cells: role of GnRH receptor concentration. Endocrinology. 2003;144:1802–1811. 57. Kirk SE, Dalkin AC, Yasin M, Haisenleder DJ, Marshall JC. Gonadotropin-­ releasing hormone pulse frequency regulates expression of pituitary follistatin messenger ribonucleic acid: a mechanism for differential gonadotrope function. Endocrinology. 1994;135:876–880. 58. Clarke IJ, Cummins JT. The temporal relationship between gonadotropin releasing hormone (GnRH) and luteinizing hormone (LH) secretion in ovariectomized ewes. Endocrinology. 1982;111:1737–1739. 59. Whitcomb RW, O’Dea LS, Finkelstein JS, Heavern DM, Crowley WF Jr. Utility of free alpha-­subunit as an alternative neuroendocrine marker of gonadotropin-­releasing hormone (GnRH) stimulation of the gonadotroph in the human: evidence from normal and GnRH-­deficient men. J Clin Endocrinol Metab. 1990;70:1654–1661. 60. Hayes FJ, McNicholl DJ, Schoenfeld D, Marsh EE, Hall JE. Free alpha-­subunit is superior to luteinizing hormone as a marker of gonadotropin-­ releasing hormone despite desensitization at fast pulse frequencies. J Clin Endocrinol Metab. 1999;84:1028–1036. 61. Clarke IJ, Burman KJ, Doughton BW, Cummins JT. Effects of constant infusion of gonadotrophin-­releasing hormone in ovariectomized ewes with hypothalamo-­ pituitary disconnection: further evidence for differential control of LH and FSH secretion and the lack of a priming effect. J Endocrinol. 1986;111:43–49. 62. Clarke I, Moore L, Veldhuis J. Intensive direct cavernous sinus sampling identifies high-­frequency, nearly random patterns of FSH secretion in ovariectomized ewes: combined appraisal by RIA and bioassay. Endocrinology. 2002;143:117–129. 63. Hall JE, Whitcomb RW, Rivier JE, Vale WW, Crowley WF Jr. Differential regulation of luteinizing hormone, follicle-­stimulating hormone, and free alpha-­subunit secretion from the gonadotrope by gonadotropin-­releasing hormone (GnRH): evidence from the use of two GnRH antagonists. J Clin Endocrinol Metab. 1990;70:328–335. 64. Gottsch ML, Cunningham MJ, Smith JT, et al. A role for kisspeptins in the regulation of gonadotropin secretion in the mouse. Endocrinology. 2004;145:4073–4077.

65. de Roux N, Genin E, Carel JC, Matsuda F, Chaussain JL, Milgrom E. Hypogonadotropic hypogonadism due to loss of function of the KiSS1-­derived peptide receptor GPR54. Proc Natl Acad Sci USA. 2003;100:10972–10976. 66. Seminara SB, Messager S, Chatzidaki EE, et al. The GPR54 gene as a regulator of puberty. N Engl J Med. 2003;349:1614–1627. 67. Topaloglu AK, Tello JA, Kotan LD, et al. Inactivating KISS1 mutation and hypogonadotropic hypogonadism. N Engl J Med. 2012;366:629–635. 68.  d’Anglemont de Tassigny X, Fagg LA, Dixon JP, et al. Hypogonadotropic hypogonadism in mice lacking a functional Kiss1 gene. Proc Natl Acad Sci USA. 2007;104:10714–10719. 69. Lapatto R, Pallais JC, Zhang D, et al. Kiss1-­ /-­mice exhibit more variable hypogonadism than Gpr54-­/-­mice. Endocrinology. 2007;148:4927–4936. 70. Teles MG, Bianco SD, Brito VN, et al. A GPR54-­activating mutation in a patient with central precocious puberty. N Engl J Med. 2008;358:709–715. 71. Silveira LG, Noel SD, Silveira-­Neto AP, et al. Mutations of the KISS1 gene in disorders of puberty. J Clin Endocrinol Metab. 2010;95:2276–2280. 72. Clarkson J, Herbison AE. Postnatal development of kisspeptin neurons in mouse hypothalamus; sexual dimorphism and projections to gonadotropin-­releasing hormone neurons. Endocrinology. 2006;147:5817–5825. 73. Yeo SH, Herbison AE. Projections of arcuate nucleus and rostral periventricular kisspeptin neurons in the adult female mouse brain. Endocrinology. 2011;152:2387–2399. 74. Kumar D, Freese M, Drexler D, Hermans-­Borgmeyer I, Marquardt A, Boehm U. Murine arcuate nucleus kisspeptin neurons communicate with GnRH neurons in utero. J Neurosci. 2014;34:3756–3766. 75. Kumar D, Periasamy V, Freese M, Voigt A, Boehm U. In utero development of kisspeptin/GnRH neural circuitry in male mice. Endocrinology. 2015;156:3084–3090. 76. Irwig MS, Fraley GS, Smith JT, et al. Kisspeptin activation of gonadotropin releasing hormone neurons and regulation of KiSS-­1 mRNA in the male rat. Neuroendocrinology. 2004;80:264–272. 77. Herbison AE, de Tassigny X, Doran J, Colledge WH. Distribution and postnatal development of Gpr54 gene expression in mouse brain and gonadotropin-­releasing hormone neurons. Endocrinology. 2010;151:312–321. 78. Smith JT, Li Q, Yap KS, et al. Kisspeptin is essential for the full preovulatory LH surge and stimulates GnRH release from the isolated ovine median eminence. Endocrinology. 2011;152:1001–1012. 79. Han SK, Gottsch ML, Lee KJ, et al. Activation of gonadotropin-­ releasing hormone neurons by kisspeptin as a neuroendocrine switch for the onset of puberty. J Neurosci. 2005;25:11349–11356. 80. Pielecka-­Fortuna J, Chu Z, Moenter SM. Kisspeptin acts directly and indirectly to increase gonadotropin-­releasing hormone neuron activity and its effects are modulated by estradiol. Endocrinology. 2008;149:1979–1986. 81. Roseweir AK, Kauffman AS, Smith JT, et al. Discovery of potent kisspeptin antagonists delineate physiological mechanisms of gonadotropin regulation. J Neurosci. 2009;29:3920–3929. 82. Shahab M, Mastronardi C, Seminara SB, Crowley WF, Ojeda SR, Plant TM. Increased hypothalamic GPR54 signaling: a potential mechanism for initiation of puberty in primates. Proc Natl Acad Sci USA. 2005;102:2129–2134. 83. Pielecka-­Fortuna J, Moenter SM. Kisspeptin increases gamma-­ aminobutyric acidergic and glutamatergic transmission directly to gonadotropin-­releasing hormone neurons in an estradiol-­dependent manner. Endocrinology. 2010;151:291–300. 84. Messager S, Chatzidaki EE, Ma D, et al. Kisspeptin directly stimulates gonadotropin-­releasing hormone release via G protein-­coupled receptor 54. Proc Natl Acad Sci USA. 2005;102:1761–1766. 85. Kirilov M, Clarkson J, Liu X, et al. Dependence of fertility on kisspeptin-­Gpr54 signaling at the GnRH neuron. Nat Commun. 2013;4:2492. 86. Novaira HJ, Sonko ML, Hoffman G, et al. Disrupted kisspeptin signaling in GnRH neurons leads to hypogonadotrophic hypogonadism. Mol Endocrinol. 2014;28:225–238. 87. Lehman MN, Coolen LM, Goodman RL. Minireview: kisspeptin/ neurokinin B/dynorphin (KNDy) cells of the arcuate nucleus: a central node in the control of gonadotropin-­releasing hormone secre-

CHAPTER 1  Neuroendocrinology of Reproduction 88. Ramaswamy S, Guerriero KA, Gibbs RB, Plant TM. Structural interactions between kisspeptin and GnRH neurons in the mediobasal hypothalamus of the male rhesus monkey (Macaca mulatta) as revealed by double immunofluorescence and confocal microscopy. Endocrinology. 2008;149:4387–4395. 89. d’Anglemont de Tassigny X, Fagg LA, Carlton MB, Colledge WH. Kisspeptin can stimulate gonadotropin-­releasing hormone (GnRH) release by a direct action at GnRH nerve terminals. Endocrinology. 2008;149:3926–3932. 90. Pallais JC, Bo-­Abbas Y, Pitteloud N, Crowley WF Jr, Seminara SB. Neuroendocrine, gonadal, placental, and obstetric phenotypes in patients with IHH and mutations in the G-­protein coupled receptor, GPR54. Mol Cell Endocrinol. 2006;254–255:70–77. 91. Rometo AM, Krajewski SJ, Voytko ML, Rance NE. Hypertrophy and increased kisspeptin gene expression in the hypothalamic infundibular nucleus of postmenopausal women and ovariectomized monkeys. J Clin Endocrinol Metab. 2007;92:2744–2750. 92. Hrabovszky E, Ciofi P, Vida B, et al. The kisspeptin system of the human hypothalamus: sexual dimorphism and relationship with gonadotropin-­releasing hormone and neurokinin B neurons. Eur J Neurosci. 2010;31:1984–1998. 93. Lehman MN, Hileman SM, Goodman RL. Neuroanatomy of the kisspeptin signaling system in mammals: comparative and developmental aspects. Adv Exp Med. 2013;784:27–62. 94. Smith JT, Cunningham MJ, Rissman EF, Clifton DK, Steiner RA. Regulation of Kiss1 gene expression in the brain of the female mouse. Endocrinology. 2005;146:3686–3692. 95. Kauffman AS, Gottsch ML, Roa J, et al. Sexual differentiation of Kiss1 gene expression in the brain of the rat. Endocrinology. 2007;148:1774–1783. 96. Homma T, Sakakibara M, Yamada S, et al. Significance of neonatal testicular sex steroids to defeminize anteroventral periventricular kisspeptin neurons and the GnRH/LH surge system in male rats. Biol Reprod. 2009;81:1216–1225. 97. Cheng G, Coolen LM, Padmanabhan V, Goodman RL, Lehman MN. The kisspeptin/neurokinin B/dynorphin (KNDy) cell population of the arcuate nucleus: sex differences and effects of prenatal testosterone in sheep. Endocrinology. 2010;151:301–311. 98. Rumpler E, Skrapits K, Takacs S, et al. Characterization of kisspeptin neurons in the human rostral hypothalamus. Neuroendocrinology. 2021;111:249–262. 99. Dhillo WS, Chaudhri OB, Thompson EL, et al. Kisspeptin-­ 54 stimulates gonadotropin release most potently during the preovulatory phase of the menstrual cycle in women. J Clin Endocrinol Metab. 2007;92:3958–3966. 100. Jayasena CN, Nijher GM, Comninos AN, et al. The effects of kisspeptin-­10 on reproductive hormone release show sexual dimorphism in humans. J Clin Endocrinol Metab. 2011;96:E1963–E1972. 101. Chan YM, Butler JP, Sidhoum VF, Pinnell NE, Seminara SB. Kisspeptin administration to women: a window into endogenous kisspeptin secretion and GnRH responsiveness across the menstrual cycle. J Clin Endocrinol Metab. 2012;97:E1458–E1467. 102. Narayanaswamy S, Jayasena CN, Ng N, et al. Subcutaneous infusion of kisspeptin-­54 stimulates gonadotrophin release in women and the response correlates with basal oestradiol levels. Clin Endocrinol. 2016;84:939–945. 103. Lippincott MF, Chan YM, Rivera Morales D, Seminara SB. Continuous kisspeptin administration in postmenopausal women: impact of estradiol on luteinizing hormone secretion. J Clin Endocrinol Metab. 2017;102:2091–2099. 104. Jayasena CN, Nijher GM, Chaudhri OB, et al. Subcutaneous injection of kisspeptin-­54 acutely stimulates gonadotropin secretion in women with hypothalamic amenorrhea, but chronic administration causes tachyphylaxis. J Clin Endocrinol Metab. 2009;94:4315–4323. 105. George JT, Anderson RA, Millar RP. Kisspeptin-­10 stimulation of gonadotrophin secretion in women is modulated by sex steroid feedback. Hum Reprod. 2012;27:3552–3559. 106. Newton CL, Anderson RC, Millar RP. Therapeutic neuroendocrine agonist and antagonist analogs of hypothalamic neuropeptides as modulators of the hypothalamic-­pituitary-­gonadal Axis. Endocr Dev. 2016;30:106–129. 107. Jayasena CN, Nijher GM, Abbara A, et al. Twice-­weekly administration of kisspeptin-­54 for 8 weeks stimulates release of reproductive hormones in women with hypothalamic amenorrhea. Clin Pharmacol Ther

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108. Jayasena CN, Abbara A, Veldhuis JD, et al. Increasing LH pulsatility in women with hypothalamic amenorrhoea using intravenous infusion of Kisspeptin-­54. J Clin Endocrinol Metab. 2014;99:E953–E961. 109. George JT, Veldhuis JD, Tena-­Sempere M, Millar RP, Anderson RA. Exploring the pathophysiology of hypogonadism in men with type 2 diabetes: kisspeptin-­10 stimulates serum testosterone and LH secretion in men with type 2 diabetes and mild biochemical hypogonadism. Clin Endocrinol. 2013;79:100–104. 110. Millar RP, Sonigo C, Anderson RA, et al. Hypothalamic-­pituitary-­ ovarian axis reactivation by kisspeptin-­ 10 in hyperprolactinemic women with chronic amenorrhea. J Endocr Soc. 2017;1:1362–1371. 111. Abbara A, Jayasena CN, Christopoulos G, et al. Efficacy of kisspeptin-­54 to trigger oocyte maturation in women at high risk of ovarian hyperstimulation syndrome (OHSS) during in vitro fertilization (IVF) therapy. J Clin Endocrinol Metab. 2015;100:3322–3331. 112. Abbara A, Clarke S, Islam R, et al. A second dose of kisspeptin-­54 improves oocyte maturation in women at high risk of ovarian hyperstimulation syndrome: a Phase 2 randomized controlled trial. Hum Reprod. 2017;32:1915–1924. 113. Regoli D, Nguyen QT, Jukic D. Neurokinin receptor subtypes characterized by biological assays. Life Sci. 1994;54:2035–2047. 114. Topaloglu AK, Reimann F, Guclu M, et al. TAC3 and TACR3 mutations in familial hypogonadotropic hypogonadism reveal a key role for Neurokinin B in the central control of reproduction. Nat Genet. 2009;41:354–358. 115. Guran T, Tolhurst G, Bereket A, et al. Hypogonadotropic hypogonadism due to a novel missense mutation in the first extracellular loop of the neurokinin B receptor. J Clin Endocrinol Metab. 2009;94:3633–3639. 116. Goodman RL, Coolen LM, Lehman MN. A role for neurokinin B in pulsatile GnRH secretion in the Ewe. Neuroendocrinology. 2014;99:18–32. 117. Grachev P, Millar RP, O’Byrne KT. The role of neurokinin B signalling in reproductive neuroendocrinology. Neuroendocrinology. 2014;99:7–17. 118. Navarro VM, Castellano JM, McConkey SM, et al. Interactions between kisspeptin and neurokinin B in the control of GnRH secretion in the female rat. Am J Physiol Endocrinol Metab. 2011;300:E202–E210. 119. Billings HJ, Connors JM, Altman SN, et al. Neurokinin B acts via the neurokinin-­ 3 receptor in the retrochiasmatic area to stimulate luteinizing hormone secretion in sheep. Endocrinology. 2010;151:3836–3846. 120. Ramaswamy S, Seminara SB, Ali B, Ciofi P, Amin NA, Plant TM. Neurokinin B stimulates GnRH release in the male monkey (Macaca mulatta) and is colocalized with kisspeptin in the arcuate nucleus. Endocrinology. 2010;151:4494–4503. 121. Krajewski SJ, Anderson MJ, Iles-­Shih L, Chen KJ, Urbanski HF, Rance NE. Morphologic evidence that neurokinin B modulates gonadotropin-­releasing hormone secretion via neurokinin 3 receptors in the rat median eminence. J Comp Neurol. 2005;489:372–386. 122. Amstalden M, Coolen LM, Hemmerle AM, et al. Neurokinin 3 receptor immunoreactivity in the septal region, preoptic area and hypothalamus of the female sheep: colocalisation in neurokinin B cells of the arcuate nucleus but not in gonadotrophin-­releasing hormone neurones. J Neuroendocrinol. 2010;22:1–12. 123. Gaskins GT, Glanowska KM, Moenter SM. Activation of neurokinin 3 receptors stimulates GnRH release in a location-­dependent but kisspeptin-­ independent manner in adult mice. Endocrinology. 2013;154:3984–3989. 124. Garcia-­Galiano D, van Ingen Schenau D, Leon S, et al. Kisspeptin signaling is indispensable for neurokinin B, but not glutamate, stimulation of gonadotropin secretion in mice. Endocrinology. 2012;153:316–328. 125. Grachev P, Li XF, Lin YS, et al. GPR54-­dependent stimulation of luteinizing hormone secretion by neurokinin B in prepubertal rats. PLoS One. 2012;7:e44344. 126. Ramaswamy S, Seminara SB, Plant TM. Evidence from the agonadal juvenile male rhesus monkey (Macaca mulatta) for the view that the action of neurokinin B to trigger gonadotropin-­releasing hormone release is upstream from the kisspeptin receptor. Neuroendocrinology. 2011;94:237–245. 127. Young J, George JT, Tello JA, et al. Kisspeptin restores pulsatile LH secretion in patients with neurokinin B signaling deficiencies: physiological, pathophysiological and therapeutic implications.

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128. Siuciak JA, McCarthy SA, Martin AN, et al. Disruption of the neurokinin-­3 receptor (NK3) in mice leads to cognitive deficits. Psychopharmacology. 2007;194:185–195. 129. Yang JJ, Caligioni CS, Chan YM, Seminara SB. Uncovering novel reproductive defects in neurokinin B receptor null mice: closing the gap between mice and men. Endocrinology. 2012;153:1498–1508. 130. de Croft S, Boehm U, Herbison AE. Neurokinin B activates arcuate kisspeptin neurons through multiple tachykinin receptors in the male mouse. Endocrinology. 2013;154:2750–2760. 131. Navarro VM, Bosch MA, Leon S, et al. The integrated hypothalamic tachykinin-­ kisspeptin system as a central coordinator for reproduction. Endocrinology. 2015;156:627–637. 132. Lehman MN, He W, Coolen LM, Levine JE, Goodman RL. Does the KNDy model for the control of gonadotropin-­releasing hormone pulses apply to monkeys and humans? Semin Reprod Med. 2019;37:71–83. 133. George JT, Kakkar R, Marshall J, et al. Neurokinin B receptor antagonism in women with polycystic ovary syndrome: a randomized, placebo-­ controlled trial. J Clin Endocrinol Metab. 2016;101:4313–4321. 134. Skorupskaite K, George JT, Veldhuis JD, Millar RP, Anderson RA. Kisspeptin and neurokinin B interactions in modulating gonadotropin secretion in women with polycystic ovary syndrome. Hum Reprod. 2020;35:1421–1431. 135. Fraser GL, Hoveyda HR, Clarke IJ, et al. The NK3 receptor antagonist ESN364 interrupts pulsatile LH secretion and moderates levels of ovarian hormones throughout the menstrual cycle. Endocrinology. 2015;156:4214–4225. 136. Fraser GL, Ramael S, Hoveyda HR, Gheyle L, Combalbert J. The NK3 receptor antagonist ESN364 suppresses sex hormones in men and women. J Clin Endocrinol Metab. 2016;101:417–426. 137. Skorupskaite K, George JT, Veldhuis JD, Millar RP, Anderson RA. Interactions between neurokinin B and kisspeptin in mediating estrogen feedback in healthy women. J Clin Endocrinol Metab. 2016;101:4628–4636. 138. Skorupskaite K, George JT, Veldhuis JD, Anderson RA. Neurokinin B regulates gonadotropin secretion, ovarian follicle growth, and the timing of ovulation in healthy women. J Clin Endocrinol Metab. 2018;103:95–104. 139. Jayasena CN, Comninos AN, Stefanopoulou E, et al. Neurokinin B administration induces hot flushes in women. Sci Rep. 2015;5:8466. 140. Prague JK, Roberts RE, Comninos AN, et al. Neurokinin 3 receptor antagonism as a novel treatment for menopausal hot flushes: a phase 2, randomised, double-­blind, placebo-­controlled trial. Lancet. 2017;389:1809–1820. 141. Skorupskaite K, George JT, Veldhuis JD, Millar RP, Anderson RA. Neurokinin 3 receptor antagonism reveals roles for neurokinin B in the regulation of gonadotropin secretion and hot flashes in postmenopausal women. Neuroendocrinology. 2018;106:148–157. 142. Depypere H, Timmerman D, Donders G, et al. Treatment of menopausal vasomotor symptoms with fezolinetant, a neurokinin 3 receptor antagonist: a phase 2a trial. J Clin Endocrinol Metab. 2019;104:5893–5905. 143. Vuong C, Van Uum SH, O’Dell LE, Lutfy K, Friedman TC. The effects of opioids and opioid analogs on animal and human endocrine systems. Endocr Rev. 2010;31:98–132. 144. Akil H, Owens C, Gutstein H, Taylor L, Curran E, Watson S. Endogenous opioids: overview and current issues. Drug Alcohol Depend. 1998;51:127–140. 145. Wehrenberg WB, Wardlaw SL, Frantz AG, Ferin M. beta-­ Endorphin in hypophyseal portal blood: variations throughout the menstrual cycle. Endocrinology. 1982;111:879–881. 146. Wardlaw SL, Wehrenberg WB, Ferin M, Antunes JL, Frantz AG. Effect of sex steroids on beta-­ endorphin in hypophyseal portal blood. J Clin Endocrinol Metab. 1982;55:877–881. 147. Gindoff PR, Jewelewicz R, Hembree W, Wardlaw S, Ferin M. Sustained effects of opioid antagonism during the normal human luteal phase. J Clin Endocrinol Metab. 1988;66:1000–1004. 148. Casper RF, Alapin-­Rubillovitz S. Progestins increase endogenous opioid peptide activity in postmenopausal women. J Clin Endocrinol Metab. 1985;60:34–36. 149. Rasmussen DD, Gambacciani M, Swartz W, Tueros VS, Yen SS. Pulsatile gonadotropin-­releasing hormone release from the human mediobasal hypothalamus in pression. Neuroendocrinology

150. Foradori CD, Goodman RL, Adams VL, Valent M, Lehman MN. Progesterone increases dynorphin a concentrations in cerebrospinal fluid and preprodynorphin messenger ribonucleic Acid levels in a subset of dynorphin neurons in the sheep. Endocrinology. 2005;146:1835–1842. 151. Wakabayashi Y, Nakada T, Murata K, et al. Neurokinin B and dynorphin A in kisspeptin neurons of the arcuate nucleus participate in generation of periodic oscillation of neural activity driving pulsatile gonadotropin-­releasing hormone secretion in the goat. J Neurosci. 2010;30:3124–3132. 152. Goodman RL, Coolen LM, Anderson GM, et al. Evidence that dynorphin plays a major role in mediating progesterone negative feedback on gonadotropin-­ releasing hormone neurons in sheep. Endocrinology. 2004;145:2959–2967. 153. Foradori CD, Coolen LM, Fitzgerald ME, Skinner DC, Goodman RL, Lehman MN. Colocalization of progesterone receptors in parvicellular dynorphin neurons of the ovine preoptic area and hypothalamus. Endocrinology. 2002;143:4366–4374. 154. Dahl SK, Amstalden M, Coolen L, Fitzgerald M, Lehman M. Dynorphin immunoreactive fibers contact GnRH neurons in the human hypothalamus. Reprod Sci. 2009;16:781–787. 155. Moore AM, Coolen LM, Porter DT, Goodman RL, Lehman MN. KNDy cells revisited. Endocrinology. 2018;159:3219–3234. 156. Burke MC, Letts PA, Krajewski SJ, Rance NE. Coexpression of dynorphin and neurokinin B immunoreactivity in the rat hypothalamus: morphologic evidence of interrelated function within the arcuate nucleus. J Comp Neurol. 2006;498:712–726. 157. Navarro VM, Gottsch ML, Chavkin C, Okamura H, Clifton DK, Steiner RA. Regulation of gonadotropin-­releasing hormone secretion by kisspeptin/dynorphin/neurokinin B neurons in the arcuate nucleus of the mouse. J Neurosci. 2009;29:11859–11866. 158. Goodman RL, Lehman MN, Smith JT, et al. Kisspeptin neurons in the arcuate nucleus of the Ewe express both dynorphin A and neurokinin B. Endocrinology. 2007;148:5752–5760. 159. Foradori CD, Amstalden M, Goodman RL, Lehman MN. Colocalisation of dynorphin a and neurokinin B immunoreactivity in the arcuate nucleus and median eminence of the sheep. J Neuroendocrinol. 2006;18:534–541. 160. Wakabayashi Y, Yamamura T, Sakamoto K, Mori Y, Okamura H. Electrophysiological and morphological evidence for synchronized GnRH pulse generator activity among Kisspeptin/neurokinin B/ dynorphin A (KNDy) neurons in goats. J Reprod Dev. 2013;59:40–48. 161. True C, Kirigiti M, Ciofi P, Grove KL, Smith MS. Characterisation of arcuate nucleus kisspeptin/neurokinin B neuronal projections and regulation during lactation in the rat. J Neuroendocrinol. 2011;23:52–64. 162. Herbison AE. The gonadotropin-­releasing hormone pulse generator. Endocrinology. 2018;159:3723–3736. 163. Hrabovszky E, Molnar CS, Sipos MT, et al. Sexual dimorphism of kisspeptin and neurokinin B immunoreactive neurons in the infundibular nucleus of aged men and women. Front Endocrinol. 2011;2:80. 164. Hrabovszky E, Sipos MT, Molnar CS, et al. Low degree of overlap between kisspeptin, neurokinin B, and dynorphin immunoreactivities in the infundibular nucleus of young male human subjects challenges the KNDy neuron concept. Endocrinology. 2012;153:4978–4989. 165. Ubuka T, Tsutsui K. Reproductive neuroendocrinology of mammalian gonadotropin-­ inhibitory hormone. Reprod Med Biol. 2019;18:225–233. 166. Tsutsui K, Ubuka T. Gonadotropin-­inhibitory hormone (GnIH): a new key neurohormone controlling reproductive physiology and behavior. Front Neuroendocrinol. 2021;61:100900. 167. Ubuka T, Morgan K, Pawson AJ, et al. Identification of human GnIH homologs, RFRP-­1 and RFRP-­3, and the cognate receptor, GPR147 in the human hypothalamic pituitary axis. PLoS One. 2009;4:e8400. 168. Ubuka T, Lai H, Kitani M, et al. Gonadotropin-­inhibitory hormone identification, cDNA cloning, and distribution in rhesus macaque brain. J Comp Neurol. 2009;517:841–855. 169. Poling MC, Quennell JH, Anderson GM, Kauffman AS. Kisspeptin neurones do not directly signal to RFRP-­3 neurones but RFRP-­3 may directly modulate a subset of hypothalamic kisspeptin cells in mice. J Neuroendocrinol. 2013;25:876–886. . RFamide-­ related pep3, a mammalian gonadotropin-inhibitory hormone ortholog,

CHAPTER 1  Neuroendocrinology of Reproduction regulates gonadotropin-­ releasing hormone neuron firing in the mouse. Endocrinology. 2009;150:2799–2804. 171. Sari IP, Rao A, Smith JT, Tilbrook AJ, Clarke IJ. Effect of RF-­ amide-­ related peptide-­ 3 on luteinizing hormone and follicle-­ stimulating hormone synthesis and secretion in ovine pituitary gonadotropes. Endocrinology. 2009;150:5549–5556. 172. Clarke IJ, Sari IP, Qi Y, et al. Potent action of RFamide-­related peptide-­3 on pituitary gonadotropes indicative of a hypophysiotropic role in the negative regulation of gonadotropin secretion. Endocrinology. 2008;149:5811–5821. 173. Clarke IJ, Smith JT, Henry BA, et al. Gonadotropin-­Inhibitory hormone is a hypothalamic peptide that provides a molecular switch between reproduction and feeding. Neuroendocrinology. 2012;95:305–316. 174. Smith JT, Shahab M, Pereira A, Pau KY, Clarke IJ. Hypothalamic expression of KISS1 and gonadotropin inhibitory hormone genes during the menstrual cycle of a non-­human primate. Biol Reprod. 2010;83:568–577. 175. Thiery JC, Pelletier J. Multiunit activity in the anterior median eminence and adjacent areas of the hypothalamus of the Ewe in relation to LH secretion. Neuroendocrinology. 1981;32:217–224. 176. Wilson RC, Kesner JS, Kaufman JM, Uemura T, Akema T, Knobil E. Central electrophysiologic correlates of pulsatile luteinizing hormone secretion in the rhesus monkey. Neuroendocrinology. 1984;39:256–260. 177. Claypool LE, Watanabe G, Terasawa E. Effects of electrical stimulation of the medial basal hypothalamus on the in vivo release of luteinizing hormone-­ releasing hormone in the prepubertal and peripubertal female monkey. Endocrinology. 1990;127:3014–3022. 178. Krey LC, Butler WR, Knobil E. Surgical disconnection of the medial basal hypothalamus and pituitary function in the rhesus monkey. I. Gonadotropin secretion. Endocrinology. 1975;96:1073–1087. 179. Martinez de la Escalera G, Choi AL, Weiner RI. Generation and synchronization of gonadotropin-­ releasing hormone (GnRH) pulses: intrinsic properties of the GT1-­1 GnRH neuronal cell line. Proc Natl Acad Sci USA. 1992;89:1852–1855. 180. Wetsel WC, Valenca MM, Merchenthaler I, et al. Intrinsic pulsatile secretory activity of immortalized luteinizing hormone-­ releasing hormone-­ secreting neurons. Proc Natl Acad Sci USA. 1992;89:4149–4153. 181. Terasawa E, Quanbeck CD, Schulz CA, Burich AJ, Luchansky LL, Claude P. A primary cell culture system of luteinizing hormone releasing hormone neurons derived from embryonic olfactory placode in the rhesus monkey. Endocrinology. 1993;133:2379–2390. 182. Funabashi T, Daikoku S, Shinohara K, Kimura F. Pulsatile gonadotropin-­releasing hormone (GnRH) secretion is an inherent function of GnRH neurons, as revealed by the culture of medial olfactory placode obtained from embryonic rats. Neuroendocrinology. 2000;71:138–144. 183. Duittoz AH, Batailler M. Pulsatile GnRH secretion from primary cultures of sheep olfactory placode explants. J Reprod Fertil. 2000;120:391–396. 184. Leranth C, Segura LM, Palkovits M, MacLusky NJ, Shanabrough M, Naftolin F. The LH-­RH-­containing neuronal network in the preoptic area of the rat: demonstration of LH-­RH-­containing nerve terminals in synaptic contact with LH-­ RH neurons. Brain Res. 1985;345:332–336. 185. Maeda K, Ohkura S, Uenoyama Y, et al. Neurobiological mechanisms underlying GnRH pulse generation by the hypothalamus. Brain Res. 2010;1364:103–115. 186. Ohkura S, Takase K, Matsuyama S, et al. Gonadotrophin-­releasing hormone pulse generator activity in the hypothalamus of the goat. J Neuroendocrinol. 2009;21:813–821. 187. Smith JT, Rao A, Pereira A, Caraty A, Millar RP, Clarke IJ. Kisspeptin is present in ovine hypophysial portal blood but does not increase during the preovulatory luteinizing hormone surge: evidence that gonadotropes are not direct targets of kisspeptin in vivo. Endocrinology. 2008;149:1951–1959. 188. Keen KL, Wegner FH, Bloom SR, Ghatei MA, Terasawa E. An increase in kisspeptin-­54 release occurs with the pubertal increase in luteinizing hormone-­releasing hormone-­1 release in the stalk-­ median eminence of female rhesus monkeys in vivo. Endocrinology. 2008;149:4151–4157. 189. Choe HK, Kim HD, Park SH, et al. Synchronous activation of gonadotropin-

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secretion by pulsatile kisspeptin stimulation. Proc Natl Acad Sci USA. 2013;110:5677–5682. 190. Clarkson J, Han SY, Piet R, et al. Definition of the hypothalamic GnRH pulse generator in mice. Proc Natl Acad Sci USA. 2017;114:E10216–E10223. 191. Ezzat A, Pereira A, Clarke IJ. Kisspeptin is a component of the pulse generator for GnRH secretion in female sheep but not the pulse generator. Endocrinology. 2015;156:1828–1837. 192. George JT, Veldhuis JD, Roseweir AK, et al. Kisspeptin-­10 is a potent stimulator of LH and increases pulse frequency in men. J Clin Endocrinol Metab. 2011;96:E1228–E1236. 193. Chan YM, Butler JP, Pinnell NE, et al. Kisspeptin resets the hypothalamic GnRH clock in men. J Clin Endocrinol Metab. 2011;96:E908–E915. 194. Jayasena CN, Comninos AN, Veldhuis JD, et al. A single injection of kisspeptin-­54 temporarily increases luteinizing hormone pulsatility in healthy women. Clin Endocrinol. 2013;79:558–563. 195. Weems PW, Witty CF, Amstalden M, Coolen LM, Goodman RL, Lehman MN. Kappa-­Opioid receptor is colocalized in GnRH and KNDy cells in the female ovine and rat brain. Endocrinology. 2016;157:2367–2379. 196. Ruka KA, Burger LL, Moenter SM. Regulation of arcuate neurons coexpressing kisspeptin, neurokinin B, and dynorphin by modulators of neurokinin 3 and kappa-­opioid receptors in adult male mice. Endocrinology. 2013;154:2761–2771. 197. Mostari P, Ieda N, Deura C, et al. dynorphin-­kappa opioid receptor signaling partly mediates estrogen negative feedback effect on LH pulses in female rats. J Reprod Dev. 2013;59:266–272. 198. Goodman RL, Hileman SM, Nestor CC, et al. Kisspeptin, neurokinin B, and dynorphin act in the arcuate nucleus to control activity of the GnRH pulse generator in ewes. Endocrinology. 2013;154:4259–4269. 199. Tenenbaum-­Rakover Y, Commenges-­Ducos M, Iovane A, Aumas C, Admoni O, de Roux N. Neuroendocrine phenotype analysis in five patients with isolated hypogonadotropic hypogonadism due to a L102P inactivating mutation of GPR54. J Clin Endocrinol Metab. 2007;92:1137–1144. 200. Mayer C, Boehm U. Female reproductive maturation in the absence of kisspeptin/GPR54 signaling. Nat Neurosci. 2011;14:704–710. 201. Semple RK, Achermann JC, Ellery J, et al. Two novel missense mutations in g protein-­coupled receptor 54 in a patient with hypogonadotropic hypogonadism. J Clin Endocrinol Metab. 2005;90:1849–1855. 202. Jakacki RI, Kelch RP, Sauder SE, Lloyd JS, Hopwood NJ, Marshall JC. Pulsatile secretion of luteinizing hormone in children. J Clin Endocrinol Metab. 1982;55:453–458. 203. McCartney CR. Maturation of sleep-­wake gonadotrophin-­releasing hormone secretion across puberty in girls: potential mechanisms and relevance to the pathogenesis of polycystic ovary syndrome. J Neuroendocrinol. 2010;22:701–709. 204. Wu FC, Butler GE, Kelnar CJ, Huhtaniemi I, Veldhuis JD. Ontogeny of pulsatile gonadotropin releasing hormone secretion from midchildhood, through puberty, to adulthood in the human male: a study using deconvolution analysis and an ultrasensitive immunofluorometric assay. J Clin Endocrinol Metab. 1996;81:1798–1805. 205. Apter D, Butzow TL, Laughlin GA, Yen SS. Gonadotropin-­releasing hormone pulse generator activity during pubertal transition in girls: pulsatile and diurnal patterns of circulating gonadotropins. J Clin Endocrinol Metab. 1993;76:940–949. 206. Abreu AP, Dauber A, Macedo DB, et al. Central precocious puberty caused by mutations in the imprinted gene MKRN3. N Engl J Med. 2013;368:2467–2475. 207. Macedo DB, Abreu AP, Reis AC, et al. Central precocious puberty that appears to be sporadic caused by paternally inherited mutations in the imprinted gene makorin ring finger 3. J Clin Endocrinol Metab. 2014;99:E1097–E1103. 208. Bessa DS, Macedo DB, Brito VN, et al. High frequency of MKRN3 mutations in male central precocious puberty previously classified as idiopathic. Neuroendocrinology. 2017;105(1):17–25. 209. Dauber A, Cunha-­Silva M, Macedo DB, et al. Paternally inherited DLK1 deletion associated with familial central precocious puberty. J Clin Endocrinol Metab. 2017;102:1557–1567. 210. Gomes LG, Cunha-­Silva M, Crespo RP, et al. DLK1 is a novel link between reproduction and metabolism. J Clin Endocrinol Metab.

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211. O’Byrne KT, Thalabard JC, Grosser PM, et al. Radiotelemetric monitoring of hypothalamic gonadotropin-­releasing hormone pulse generator activity throughout the menstrual cycle of the rhesus monkey. Endocrinology. 1991;129:1207–1214. 212. Reame N, Sauder SE, Kelch RP, Marshall JC. Pulsatile gonadotropin secretion during the human menstrual cycle: evidence for altered frequency of gonadotropin-­releasing hormone secretion. J Clin Endocrinol Metab. 1984;59:328–337. 213. Adams JM, Taylor AE, Schoenfeld DA, Crowley WF Jr, Hall JE. The midcycle gonadotropin surge in normal women occurs in the face of an unchanging gonadotropin-­releasing hormone pulse frequency. J Clin Endocrinol Metab. 1994;79:858–864. 214. Lam NY, Ferin M. Is the decrease in the hypophysiotropic signal frequency normally observed during the luteal phase important for menstrual cyclicity in the primate? Endocrinology. 1987;120:2044–2049. 215. Welt CK, Martin KA, Taylor AE, et al. Frequency modulation of follicle-­stimulating hormone (FSH) during the luteal-­follicular transition: evidence for FSH control of inhibin B in normal women. J Clin Endocrinol Metab. 1997;82:2645–2652. 216. Dierschke DJ, Bhattacharya AN, Atkinson LE, Knobil E. Circhoral oscillations of plasma LH levels in the ovariectomized rhesus monkey. Endocrinology. 1970;87:850–853. 217. Rossmanith WG, Liu CH, Laughlin GA, Mortola JF, Suh BY, Yen SS. Relative changes in LH pulsatility during the menstrual cycle: using data from hypogonadal women as a reference point. Clin Endocrinol. 1990;32:647–660. 218. Gill S, Lavoie HB, Bo-­Abbas Y, Hall JE. Negative feedback effects of gonadal steroids are preserved with aging in postmenopausal women. J Clin Endocrinol Metab. 2002;87:2297–2302. 219. Filicori M, Santoro N, Merriam GR, Crowley WF Jr. Characterization of the physiological pattern of episodic gonadotropin secretion throughout the human menstrual cycle. J Clin Endocrinol Metab. 1986;62:1136–1144. 220. O’Dea LS, Finkelstein JS, Schoenfeld DA, Butler JP, Crowley WF Jr. Interpulse interval of GnRH stimulation independently modulates LH secretion. Am J Physiol. 1989;256:E510–E515. 221. Piotrowska K, Wang C, Swerdloff RS, Liu PY. Male hormonal contraception: hope and promise. Lancet Diabetes Endocrinol. 2017;5(3):214–223. 222. Hall JE. Neuroendocrine changes with reproductive aging in women. Semin Reprod Med. 2007;25:344–351. 223. Morishima A, Grumbach MM, Simpson ER, Fisher C, Qin K. Aromatase deficiency in male and female siblings caused by a novel mutation and the physiological role of estrogens. J Clin Endocrinol Metab. 1995;80:3689–3698. 224. Herbison AE. Multimodal influence of estrogen upon gonadotropin-­ releasing hormone neurons. Endocr Rev. 1998;19:302–330. 225. Caraty A, Locatelli A, Martin GB. Biphasic response in the secretion of gonadotrophin-­releasing hormone in ovariectomized ewes injected with oestradiol. J Endocrinol. 1989;123:375–382. 226. Chongthammakun S, Terasawa E. Negative feedback effects of estrogen on luteinizing hormone-­releasing hormone release occur in pubertal, but not prepubertal, ovariectomized female rhesus monkeys. Endocrinology. 1993;132:735–743. 227. Gill S, Sharpless JL, Rado K, Hall JE. Evidence that GnRH decreases with gonadal steroid feedback but increases with age in postmenopausal women. J Clin Endocrinol Metab. 2002;87:2290–2296. 228. Evans NP, Dahl GE, Glover BH, Karsch FJ. Central regulation of pulsatile gonadotropin-­ releasing hormone (GnRH) secretion by estradiol during the period leading up to the preovulatory GnRH surge in the Ewe. Endocrinology. 1994;134:1806–1811. 229. Kesner JS, Wilson RC, Kaufman JM, et al. Unexpected responses of the hypothalamic gonadotropin-­releasing hormone “pulse generator” to physiological estradiol inputs in the absence of the ovary. Proc Natl Acad Sci USA. 1987;84:8745–8749. 230. Cagnacci A, Melis GB, Paoletti AM, et al. Influence of oestradiol and progesterone on pulsatile LH secretion in postmenopausal women. Clin Endocrinol. 1989;31:541–550. 231. Nakai Y, Plant TM, Hess DL, Keogh EJ, Knobil E. On the sites of the negative and positive feedback actions of estradiol in the control of gonadotropin secretion in the rhesus monkey. Endocrinology. 1978;102:1008–1014. 232. Clarke IJ, Cummins JT. Direct pituitary effects of estrogen and progesterone on gonadotropin secretion in the ovariectomized Ewe. Neuroendocrinology. 1984;39:267–274.

233. Taylor AE, Whitney H, Hall JE, Martin K, Crowley WF Jr. Midcycle levels of sex steroids are sufficient to recreate the follicle-­stimulating hormone but not the luteinizing hormone midcycle surge: evidence for the contribution of other ovarian factors to the surge in normal women. J Clin Endocrinol Metab. 1995;80:1541–1547. 234. Hall JE, Schoenfeld DA, Martin KA, Crowley WF Jr. Hypothalamic gonadotropin-­releasing hormone secretion and follicle-­stimulating hormone dynamics during the luteal-­follicular transition. J Clin Endocrinol Metab. 1992;74:600–607. 235. Soules MR, Steiner RA, Clifton DK, Cohen NL, Aksel S, Bremner WJ. Progesterone modulation of pulsatile luteinizing hormone secretion in normal women. J Clin Endocrinol Metab. 1984;58:378–383. 236. Karsch FJ, Weick RF, Hotchkiss J, Dierschke DJ, Knobil E. An analysis of the negative feedback control of gonadotropin secretion utilizing chronic implantation of ovarian steroids in ovariectomized rhesus monkeys. Endocrinology. 1973;93:478–486. 237. Nippoldt TB, Reame NE, Kelch RP, Marshall JC. The roles of estradiol and progesterone in decreasing luteinizing hormone pulse frequency in the luteal phase of the menstrual cycle. J Clin Endocrinol Metab. 1989;69:67–76. 238. MacLusky NJ, McEwen BS. Oestrogen modulates progestin receptor concentrations in some rat brain regions but not in others. Nature. 1978;274:276–278. 239. Romano GJ, Krust A, Pfaff DW. Expression and estrogen regulation of progesterone receptor mRNA in neurons of the mediobasal hypothalamus: an in situ hybridization study. Mol Endocrinol. 1989;3:1295–1300. 240. Pielecka J, Quaynor SD, Moenter SM. Androgens increase gonadotropin-­releasing hormone neuron firing activity in females and interfere with progesterone negative feedback. Endocrinology. 2006;147:1474–1479. 241. Pastor CL, Griffin-­Korf ML, Aloi JA, Evans WS, Marshall JC. Polycystic ovary syndrome: evidence for reduced sensitivity of the gonadotropin-­ releasing hormone pulse generator to inhibition by estradiol and progesterone. J Clin Endocrinol Metab. 1998;83:582–590. 242. Eagleson CA, Gingrich MB, Pastor CL, et al. Polycystic ovarian syndrome: evidence that flutamide restores sensitivity of the gonadotropin-­ releasing hormone pulse generator to inhibition by estradiol and progesterone. J Clin Endocrinol Metab. 2000;85:4047–4052. 243. Foecking EM, Levine JE. Effects of experimental hyperandrogenemia on the female rat reproductive axis: suppression of progesterone-­receptor messenger RNA expression in the brain and blockade of luteinizing hormone surges. Gend Med. 2005;2:155–165. 244. Karsch FJ, Weick RF, Butler WR, et al. Induced LH surges in the rhesus monkey: strength-­duration characteristics of the estrogen stimulus. Endocrinology. 1973;92:1740–1747. 245. Liu JH, Yen SS. Induction of midcycle gonadotropin surge by ovarian steroids in women: a critical evaluation. J Clin Endocrinol Metab. 1983;57:797–802. 246. Wang CF, Lasley BL, Lein A, Yen SS. The functional changes of the pituitary gonadotrophs during the menstrual cycle. J Clin Endocrinol Metab. 1976;42:718–728. 247. Hall JE, Martin KA, Whitney HA, Landy H, Crowley WF Jr. Potential for fertility with replacement of hypothalamic gonadotropin-­releasing hormone in long term female survivors of cranial tumors. J Clin Endocrinol Metab. 1994;79:1166–1172. 248. Sarkar DK, Chiappa SA, Fink G, Sherwood NM. Gonadotropin-­ releasing hormone surge in pro-­ oestrous rats. Nature. 1976;264:461–463. 249. Moenter SM, Caraty A, Karsch FJ. The estradiol-­induced surge of gonadotropin-­ releasing hormone in the Ewe. Endocrinology. 1990;127:1375–1384. 250. Levine JE, Norman RL, Gliessman PM, Oyama TT, Bangsberg DR, Spies HG. In vivo gonadotropin-­releasing hormone release and serum luteinizing hormone measurements in ovariectomized, estrogen-­treated rhesus macaques. Endocrinology. 1985;117:711–721. 251. Pau KY, Berria M, Hess DL, Spies HG. Preovulatory gonadotropin-­ releasing hormone surge in ovarian-­ intact rhesus macaques. Endocrinology. 1993;133:1650–1656. 252. Knobil E, Plant TM, Wildt L, Belchetz PE, Marshall G. Control of the rhesus monkey menstrual cycle: permissive role of hypothalamic gonadotropin-­releasing hormone. Science. 1980;207:1371–1373. , Schoenfeld DA, Crowley . Decreased release of gonadotropin-releasing hormone

CHAPTER 1  Neuroendocrinology of Reproduction during the preovulatory midcycle luteinizing hormone surge in normal women. Proc Natl Acad Sci USA. 1994;91:6894–6898. 254. Martin KA, Welt CK, Taylor AE, Smith JA, Crowley WF Jr, Hall JE. Is GnRH reduced at the midcycle surge in the human? Evidence from a GnRH-­ deficient model. Neuroendocrinology. 1998;67:363–369. 255. Ottowitz WE, Dougherty DD, Fischman AJ, Hall JE. [18F]2-­fluoro-­ 2-­deoxy-­D-­glucose positron emission tomography demonstration of estrogen negative and positive feedback on luteinizing hormone secretion in women. J Clin Endocrinol Metab. 2008;93:3208–3214. 256. Kolp LA, Pavlou SN, Urban RJ, Rivier JC, Vale WW, Veldhuis JD. Abrogation by a potent gonadotropin-­releasing hormone antagonist of the estrogen/progesterone-­stimulated surge-­like release of luteinizing hormone and follicle-­stimulating hormone in postmenopausal women. J Clin Endocrinol Metab. 1992;75:993–997. 257. Dubourdieu S, Charbonnel B, D’Acremont MF, Carreau S, Spitz IM, Bouchard P. Effect of administration of a gonadotropin-­ releasing hormone (GnRH) antagonist (Nal-­Glu) during the periovulatory period: the luteinizing hormone surge requires secretion of GnRH. J Clin Endocrinol Metab. 1994;78:343–347. 258. Nippoldt TB, Khoury S, Barkan A, Kelch RP, Marshall JC. Gonadotrophin responses to GnRH pulses in hypogonadotrophic hypogonadism: LH responsiveness is maintained in the presence of luteal phase concentrations of oestrogen and progesterone. Clin Endocrinol. 1987;26:293–301. 259. Couzinet B, Brailly S, Bouchard P, Schaison G. Progesterone stimulates luteinizing hormone secretion by acting directly on the pituitary. J Clin Endocrinol Metab. 1992;74:374–378. 260. Schaison G, George M, Lestrat N, Reinberg A, Baulieu EE. Effects of the antiprogesterone steroid RU 486 during midluteal phase in normal women. J Clin Endocrinol Metab. 1985;61:484–489. 261. Garzo VG, Liu J, Ulmann A, Baulieu E, Yen SS. Effects of an antiprogesterone (RU486) on the hypothalamic-­hypophyseal-­ovarian-­ endometrial axis during the luteal phase of the menstrual cycle. J Clin Endocrinol Metab. 1988;66:508–517. 262. March CM, Goebelsmann U, Nakamura RM, Mishell DR Jr. Roles of estradiol and progesterone in eliciting the midcycle luteinizing hormone and follicle-­stimulating hormone surges. J Clin Endocrinol Metab. 1979;49:507–513. 263. Chang RJ, Jaffe RB. Progesterone effects on gonadotropin release in women pretreated with estradiol. J Clin Endocrinol Metab. 1978;47:119–125. 264. Hoff JD, Quigley ME, Yen SS. Hormonal dynamics at midcycle: a reevaluation. J Clin Endocrinol Metab. 1983;57:792–796. 265. Batista MC, Cartledge TP, Zellmer AW, Nieman LK, Loriaux DL, Merriam GR. The antiprogestin RU486 delays the midcycle gonadotropin surge and ovulation in gonadotropin-­releasing hormone-­ induced cycles. Fertil Steril. 1994;62:28–34. 266. Yen SS, Tsai CC. Acute gonadotropin release induced by exogenous estradiol during the mid-­follicular phase of the menstrual cycle. J Clin Endocrinol Metab. 1972;34:298–305. 267. Karsch FJ, Dierschke DJ, Knobil E. Sexual differentiation of pituitary function: apparent difference between primates and rodents. Science. 1973;179:484–486. 268. Norman RL, Spies HG. Cyclic ovarian function in a male macaque: additional evidence for a lack of sexual differentiation in the physiological mechanisms that regulate the cyclic release of gonadotropins in primates. Endocrinology. 1986;118:2608–2610. 269. Barbarino A, de Marinis L, Lafuenti G, Muscatello P, Matteucci BR. Presence of positive feedback between oestrogen and LH in patients with Klinefelter’s syndrome, and Sertoli-­cell-­only syndrome. Clin Endocrinol. 1979;10:235–242. 270. Stearns EL, Winter JS, Faiman C. Positive feedback effect of progestin upon serum gonadotropins in estrogen-­primed castrate men. J Clin Endocrinol Metab. 1973;37:635–638. 271. Plant TM, Hess DL, Hotchkiss J, Knobil E. Testosterone and the control of gonadotropin secretion in the male rhesus monkey (Macaca mulatta). Endocrinology. 1978;103:535–541. 272. Matsumoto AM, Bremner WJ. Modulation of pulsatile gonadotropin secretion by testosterone in man. J Clin Endocrinol Metab. 1984;58:609–614. 273. Schnorr JA, Bray MJ, Veldhuis JD. Aromatization mediates testosterone’s short-­term feedback restraint of 24-­hour endogenously driven and acute exogenous gonadotropin-­releasing hormone-­stimulated luteinizing hormone and follicleyoung men. J Clin Endocrinol Metab

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274. Pitteloud N, Dwyer AA, DeCruz S, et al. Inhibition of luteinizing hormone secretion by testosterone in men requires aromatization for its pituitary but not its hypothalamic effects: evidence from the tandem study of normal and gonadotropin-­ releasing hormone-­ deficient men. J Clin Endocrinol Metab. 2008;93:784–791. 275. Boyar RM, Moore RJ, Rosner W, et al. Studies of gonadotropin-­ gonadal dynamics in patients with androgen insensitivity. J Clin Endocrinol Metab. 1978;47:1116–1122. 276. Gooren L, Spinder T, Spijkstra JJ, et al. Sex steroids and pulsatile luteinizing hormone release in men. studies in estrogen-­treated agonadal subjects and eugonadal subjects treated with a novel nonsteroidal antiandrogen. J Clin Endocrinol Metab. 1987;64:763–770. 277. Urban RJ, Davis MR, Rogol AD, Johnson ML, Veldhuis JD. Acute androgen receptor blockade increases luteinizing hormone secretory activity in men. J Clin Endocrinol Metab. 1988;67:1149–1155. 278. Santen RJ. Is aromatization of testosterone to estradiol required for inhibition of luteinizing hormone secretion in men? J Clin Investig. 1975;56:1555–1563. 279. Kuhn JM, Rieu M, Laudat MH, et al. Effects of 10 days administration of percutaneous dihydrotestosterone on the pituitary-­testicular axis in normal men. J Clin Endocrinol Metab. 1984;58:231–235. 280. Veldhuis JD, Rogol AD, Samojlik E, Ertel NH. Role of endogenous opiates in the expression of negative feedback actions of androgen and estrogen on pulsatile properties of luteinizing hormone secretion in man. J Clin Investig. 1984;74:47–55. 281. Veldhuis JD, Dufau ML. Estradiol modulates the pulsatile secretion of biologically active luteinizing hormone in man. J Clin Investig. 1987;80:631–638. 282. Finkelstein JS, O’Dea LS, Whitcomb RW, Crowley WF Jr. Sex steroid control of gonadotropin secretion in the human male. II. Effects of estradiol administration in normal and gonadotropin-­releasing hormone-­deficient men. J Clin Endocrinol Metab. 1991;73:621–628. 283. Veldhuis JD, Urban RJ, Dufau ML. Evidence that androgen negative feedback regulates hypothalamic gonadotropin-­ releasing hormone impulse strength and the burst-­like secretion of biologically active luteinizing hormone in men. J Clin Endocrinol Metab. 1992;74:1227–1235. 284. Hayes FJ, Seminara SB, Decruz S, Boepple PA, Crowley WF Jr. Aromatase inhibition in the human male reveals a hypothalamic site of estrogen feedback. J Clin Endocrinol Metab. 2000;85:3027–3035. 285. Boyar RM, Perlow M, Kapen S, Lefkowitz G, Weitzman E, Hellman L. The effect of clomiphene citrate on the 24-­hour LH secretory pattern in normal men. J Clin Endocrinol Metab. 1973;36:561–567. 286. Santen RJ, Ruby EB. Enhanced frequency and magnitude of episodic luteinizing hormone-­releasing hormone discharge as a hypothalamic mechanism for increased luteinizing hormone secretion. J Clin Endocrinol Metab. 1979;48:315–319. 287. Winters SJ, Troen P. Evidence for a role of endogenous estrogen in the hypothalamic control of gonadotropin secretion in men. J Clin Endocrinol Metab. 1985;61:842–845. 288. Rochira V, Zirilli L, Genazzani AD, et al. Hypothalamic-­pituitary-­ gonadal axis in two men with aromatase deficiency: evidence that circulating estrogens are required at the hypothalamic level for the integrity of gonadotropin negative feedback. Eur J Endocrinol. 2006;155:513–522. 289. Bagatell CJ, Dahl KD, Bremner WJ. The direct pituitary effect of testosterone to inhibit gonadotropin secretion in men is partially mediated by aromatization to estradiol. J Androl. 1994;15:15–21. 290. Smith JT, Dungan HM, Stoll EA, et al. Differential regulation of KiSS-­1 mRNA expression by sex steroids in the brain of the male mouse. Endocrinology. 2005;146:2976–2984. 291. Rometo AM, Rance NE. Changes in prodynorphin gene expression and neuronal morphology in the hypothalamus of postmenopausal women. J Neuroendocrinol. 2008;20:1376–1381. 292. Molnar CS, Vida B, Sipos MT, et al. Morphological evidence for enhanced kisspeptin and neurokinin B signaling in the infundibular nucleus of the aging man. Endocrinology. 2012;153:5428–5439. 293. Clarkson J, d’Anglemont de Tassigny X, Moreno AS, Colledge WH, Herbison AE. Kisspeptin-­GPR54 signaling is essential for preovulatory gonadotropin-­ releasing hormone neuron activation and the luteinizing hormone surge. J Neurosci. 2008;28:8691–8697. 294. Pineda R, Garcia-­Galiano D, Roseweir A, et al. Critical roles of kisspeptins in female puberty and preovulatory gonadotropin surges as revealed by a novel antagonist. Endocrinology. 2010;151:722–730. al. Involvement of central metastin in the regulation of preovulatory luteinizing

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PART I  The Fundamentals of Reproduction

hormone surge and estrous cyclicity in female rats. Endocrinology. 2005;146:4431–4436. 296. Goodman RL. The site of the positive feedback action of estradiol in the rat. Endocrinology. 1978;102:151–159. 297. Halasz B, Gorski RA. Gonadotrophic hormone secretion in female rats after partial or total interruption of neural afferents to the medial basal hypothalamus. Endocrinology. 1967;80:608–622. 298. Plant TM, Moossy J, Hess DL, Nakai Y, McCormack JT, Knobil E. Further studies on the effects of lesions in the rostral hypothalamus on gonadotropin secretion in the female rhesus monkey (Macaca mulatta). Endocrinology. 1979;105:465–473. 299. Norman RL, Resko JA, Spies HG. The anterior hypothalamus: how it affects gonadotropin secretion in the rhesus monkey. Endocrinology. 1976;99:59–71. 300. Estrada KM, Clay CM, Pompolo S, Smith JT, Clarke IJ. Elevated KiSS-­1 expression in the arcuate nucleus prior to the cyclic preovulatory gonadotrophin-­releasing hormone/lutenising hormone surge in the Ewe suggests a stimulatory role for kisspeptin in oestrogen-­ positive feedback. J Neuroendocrinol. 2006;18:806–809. 301. Smith JT, Li Q, Pereira A, Clarke IJ. Kisspeptin neurons in the ovine arcuate nucleus and preoptic area are involved in the preovulatory luteinizing hormone surge. Endocrinology. 2009;150:5530–5538. 302. Boepple PA, Hayes FJ, Dwyer AA, et al. Relative roles of inhibin B and sex steroids in the negative feedback regulation of follicle-­ stimulating hormone in men across the full spectrum of seminiferous epithelium function. J Clin Endocrinol Metab. 2008;93:1809–1814. 303. Dalkin AC, Haisenleder DJ, Gilrain JT, Aylor K, Yasin M, Marshall JC. Gonadotropin-­releasing hormone regulation of gonadotropin subunit gene expression in female rats: actions on follicle-­stimulating hormone beta messenger ribonucleic acid (mRNA) involve differential expression of pituitary activin (beta-­B) and follistatin mRNAs. Endocrinology. 1999;140:903–908. 304. World Health Organization. Human Energy Requirements. Food and Nutrition Technical Report Series 1. Rome, Food and Agriculture Organization of the United Nations; 2004:53–62. 305. Reame NE, Sauder SE, Case GD, Kelch RP, Marshall JC. Pulsatile gonadotropin secretion in women with hypothalamic amenorrhea: evidence that reduced frequency of gonadotropin-­releasing hormone secretion is the mechanism of persistent anovulation. J Clin Endocrinol Metab. 1985;61:851–858. 306. Perkins RB, Hall JE, Martin KA. Neuroendocrine abnormalities in hypothalamic amenorrhea: spectrum, stability, and response to neurotransmitter modulation. J Clin Endocrinol Metab. 1999;84:1905–1911. 307. Berga SL, Mortola JF, Girton L, et al. Neuroendocrine aberrations in women with functional hypothalamic amenorrhea. J Clin Endocrinol Metab. 1989;68:301–308. 308. Frisch RE. The right weight: body fat, menarche, and fertility. Nutrition. 1996;12:452–453. 309. Redman LM, Loucks AB. Menstrual disorders in athletes. Sports Med. 2005;35:747–755. 310. Sterling WM, Golden NH, Jacobson MS, Ornstein RM, Hertz SM. Metabolic assessment of menstruating and nonmenstruating normal weight adolescents. Int J Eat Disord. 2009;42:658–663. 311. Di Carlo C, Palomba S, De Fazio M, Gianturco M, Armellino M, Nappi C. Hypogonadotropic hypogonadism in obese women after biliopancreatic diversion. Fertil Steril. 1999;72:905–909. 312. Loucks AB, Thuma JR. Luteinizing hormone pulsatility is disrupted at a threshold of energy availability in regularly menstruating women. J Clin Endocrinol Metab. 2003;88:297–311. 313. Lane MA, Black A, Handy AM, et al. Energy restriction does not alter bone mineral metabolism or reproductive cycling and hormones in female rhesus monkeys. J Nutr. 2001;131:820–827. 314. Myerson M, Gutin B, Warren MP, et al. Resting metabolic rate and energy balance in amenorrheic and eumenorrheic runners. Med Sci Sports Exerc. 1991;23:15–22. 315. Loucks AB, Verdun M, Heath EM. Low energy availability, not stress of exercise, alters LH pulsatility in exercising women. J Appl Physiol. 1998;84:37–46. 316. Williams NI, Caston-­Balderrama AL, Helmreich DL, Parfitt DB, Nosbisch C, Cameron JL. Longitudinal changes in reproductive hormones and menstrual cyclicity in cynomolgus monkeys during strenuous exercise training: abrupt transition to exercise-­induced amenorrhea. Endocrinology

317. Williams NI, Helmreich DL, Parfitt DB, Caston-­Balderrama A, Cameron JL. Evidence for a causal role of low energy availability in the induction of menstrual cycle disturbances during strenuous exercise training. J Clin Endocrinol Metab. 2001;86:5184–5193. 318. Misra M, Klibanski A. Neuroendocrine consequences of anorexia nervosa in adolescents. Endocr Dev. 2010;17:197–214. 319. Farooqi IS, Jebb SA, Langmack G, et al. Effects of recombinant leptin therapy in a child with congenital leptin deficiency. N Engl J Med. 1999;341:879–884. 320. Farooqi IS, Wangensteen T, Collins S, et al. Clinical and molecular genetic spectrum of congenital deficiency of the leptin receptor. N Engl J Med. 2007;356:237–247. 321. Miller KK, Parulekar MS, Schoenfeld E, et al. Decreased leptin levels in normal weight women with hypothalamic amenorrhea: the effects of body composition and nutritional intake. J Clin Endocrinol Metab. 1998;83:2309–2312. 322. Welt CK, Chan JL, Bullen J, et al. Recombinant human leptin in women with hypothalamic amenorrhea. N Engl J Med. 2004;351:987–997. 323. Chou SH, Chamberland JP, Liu X, et al. Leptin is an effective treatment for hypothalamic amenorrhea. Proc Natl Acad Sci USA. 2011;108:6585–6590. 324. Bouzoni E, Perakakis N, Mantzoros CS. Circulating profile of Activin-­ Follistatin-­ Inhibin Axis in women with hypothalamic amenorrhea in response to leptin treatment. Metab Clin Exp. 2020;113:154392. 325. Boden G, Chen X, Mozzoli M, Ryan I. Effect of fasting on serum leptin in normal human subjects. J Clin Endocrinol Metab. 1996;81:3419–3423. 326. Donato J Jr, Cravo RM, Frazao R, et al. Leptin’s effect on puberty in mice is relayed by the ventral premammillary nucleus and does not require signaling in Kiss1 neurons. J Clin Investig. 2011;121:355–368. 327. Wade GN, Jones JE. Neuroendocrinology of nutritional infertility. Am J Physiol Regul Integr Comp Physiol. 2004;287:R1277–R1296. 328. De Bond JA, Smith JT. Kisspeptin and energy balance in reproduction. Reproduction. 2014;147:R53–R63. 329. Navarro VM, Kaiser UB. Metabolic influences on neuroendocrine regulation of reproduction. Curr Opin Endocrinol Diabetes Obes. 2013;20:335–341. 330. Smith JT, Acohido BV, Clifton DK, Steiner RA. KiSS-­1 neurones are direct targets for leptin in the ob/ob mouse. J Neuroendocrinol. 2006;18:298–303. 331. Quigley ME, Sheehan KL, Casper RF, Yen SS. Evidence for increased dopaminergic and opioid activity in patients with hypothalamic hypogonadotropic amenorrhea. J Clin Endocrinol Metab. 1980;50:949–954. 332. Khoury SA, Reame NE, Kelch RP, Marshall JC. Diurnal patterns of pulsatile luteinizing hormone secretion in hypothalamic amenorrhea: reproducibility and responses to opiate blockade and an alpha 2-­adrenergic agonist. J Clin Endocrinol Metab. 1987;64:755–762. 333. Wildt L, Leyendecker G. Induction of ovulation by the chronic administration of naltrexone in hypothalamic amenorrhea. J Clin Endocrinol Metab. 1987;64:1334–1335. 334. Friedl KE, Moore RJ, Hoyt RW, Marchitelli LJ, Martinez-­ Lopez LE, Askew EW. Endocrine markers of semistarvation in healthy lean men in a multistressor environment. J Appl Physiol. 2000;88:1820–1830. 335. Misra M, Katzman DK, Cord J, et al. Percentage extremity fat, but not percentage trunk fat, is lower in adolescent boys with anorexia nervosa than in healthy adolescents. Am J Clin Nutr. 2008;88:1478–1484. 336. Dwyer AA, Chavan NR, Lewkowitz-­Shpuntoff H, et al. Functional hypogonadotropic hypogonadism in men: underlying neuroendocrine mechanisms and natural history. J Clin Endocrinol Metab. 2019;104:3403–3414. 337. Chan JL, Heist K, DePaoli AM, Veldhuis JD, Mantzoros CS. The role of falling leptin levels in the neuroendocrine and metabolic adaptation to short-­term starvation in healthy men. J Clin Investig. 2003;111:1409–1421. 338. Woolf PD, Hamill RW, McDonald JV, Lee LA, Kelly M. Transient hypogonadotropic hypogonadism caused by critical illness. J Clin Endocrinol Metab. 1985;60:444–450. 339. Spratt DI, Cox P, Orav J, Moloney J, Bigos T. Reproductive axis suppression in acute illness is related to disease severity. J Clin

CHAPTER 1  Neuroendocrinology of Reproduction 340. Williams CL, Nishihara M, Thalabard JC, Grosser PM, Hotchkiss J, Knobil E. Corticotropin-­ releasing factor and gonadotropin-­ releasing hormone pulse generator activity in the rhesus monkey. Electrophysiological studies. Neuroendocrinology. 1990;52:133–137. 341. Rivier C, Rivier J, Vale W. Stress-­induced inhibition of reproductive functions: role of endogenous corticotropin-­releasing factor. Science. 1986;231:607–609. 342. Gindoff PR, Ferin M. Endogenous opioid peptides modulate the effect of corticotropin-­releasing factor on gonadotropin release in the primate. Endocrinology. 1987;121:837–842. 343. Loucks AB, Mortola JF, Girton L, Yen SS. Alterations in the hypothalamic-­ pituitary-­ ovarian and the hypothalamic-­ pituitary-­adrenal axes in athletic women. J Clin Endocrinol Metab. 1989;68:402–411. 344. Giles DE, Berga SL. Cognitive and psychiatric correlates of functional hypothalamic amenorrhea: a controlled comparison. Fertil Steril. 1993;60:486–492. 345. Marcus MD, Loucks TL, Berga SL. Psychological correlates of functional hypothalamic amenorrhea. Fertil Steril. 2001;76:310–316. 346. Berga SL, Marcus MD, Loucks TL, Hlastala S, Ringham R, Krohn MA. Recovery of ovarian activity in women with functional hypothalamic amenorrhea who were treated with cognitive behavior therapy. Fertil Steril. 2003;80:976–981. 347. Tschugguel W, Berga SL. Treatment of functional hypothalamic amenorrhea with hypnotherapy. Fertil Steril. 2003;80:982–985. 348. Williams NI, Berga SL, Cameron JL. Synergism between psychosocial and metabolic stressors: impact on reproductive function in cynomolgus monkeys. Am J Physiol Endocrinol Metab. 2007;293:E270–E276. 349. Caronia LM, Martin C, Welt CK, et al. A genetic basis for functional hypothalamic amenorrhea. N Engl J Med. 2011;364:215–225. 350. Delaney A, Burkholder AB, Lavender CA, et al. Increased burden of rare sequence variants in GnRH-­ associated genes in women with hypothalamic amenorrhea. J Clin Endocrinol Metab. 2021;106:e1441–e1452. 351. Van der Wijden C, Manion C. Lactational amenorrhoea method for family planning. Cochrane Database Syst Rev. 2015:CD001329. 352. Thapa S, Short RV, Potts M. Breast feeding, birth spacing and their effects on child survival. Nature. 1988;335:679–682. 353. Tay CC, Glasier AF, McNeilly AS. The 24 h pattern of pulsatile luteinizing hormone, follicle stimulating hormone and prolactin release during the first 8 weeks of lactational amenorrhoea in breastfeeding women. Hum Reprod. 1992;7:951–958. 354. Ordog T, Chen MD, O’Byrne KT, et al. On the mechanism of lactational anovulation in the rhesus monkey. Am J Physiol. 1998;274:E665–E676. 355. Zinaman MJ, Cartledge T, Tomai T, Tippett P, Merriam GR. Pulsatile GnRH stimulates normal cyclic ovarian function in amenorrheic lactating postpartum women. J Clin Endocrinol Metab. 1995;80:2088–2093. 356. Picciano MF. Nutrient composition of human milk. Pediatr Clin North Am. 2001;48:53–67. 357. Smith MS, True C, Grove KL. The neuroendocrine basis of lactation-­ induced suppression of GnRH: role of kisspeptin and leptin. Brain Res. 2010;1364:139–152. 358. Xu J, Kirigiti MA, Cowley MA, Grove KL, Smith MS. Suppression of basal spontaneous gonadotropin-­ releasing hormone neuronal activity during lactation: role of inhibitory effects of neuropeptide Y. Endocrinology. 2009;150:333–340. 359. Yamada S, Uenoyama Y, Kinoshita M, et al. Inhibition of metastin (kisspeptin-­ 54)-­ GPR54 signaling in the arcuate nucleus-­ median eminence region during lactation in rats. Endocrinology. 2007;148:2226–2232. 360. Cook CB, Nippoldt TB, Kletter GB, Kelch RP, Marshall JC. Naloxone increases the frequency of pulsatile luteinizing hormone secretion in women with hyperprolactinemia. J Clin Endocrinol Metab. 1991;73:1099–1105. 361. Kokay IC, Petersen SL, Grattan DR. Identification of prolactin-­ sensitive GABA and kisspeptin neurons in regions of the rat hypothalamus involved in the control of fertility. Endocrinology. 2011;152:526–535. 362. Sonigo C, Bouilly J, Carre N, et al. Hyperprolactinemia-­induced ovarian acyclicity is reversed by kisspeptin administration. J Clin Investig. 2012;122:3791–3795.

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363. Araujo-­Lopes R, Crampton JR, Aquino NS, et al. Prolactin regulates kisspeptin neurons in the arcuate nucleus to suppress LH secretion in female rats. Endocrinology. 2014;155:1010–1020. 364. Brown RSE, Khant Aung Z, Phillipps HR, et al. Acute suppression of LH secretion by prolactin in female mice is mediated by kisspeptin neurons in the arcuate nucleus. Endocrinology. 2019;160:1323–1332. 365. Urbanski HF. Role of circadian neuroendocrine rhythms in the control of behavior and physiology. Neuroendocrinology. 2011;93:211–222. 366. Legan SJ, Karsch FJ. A daily signal for the LH surge in the rat. Endocrinology. 1975;96:57–62. 367. Kerdelhue B, Brown S, Lenoir V, et al. Timing of initiation of the preovulatory luteinizing hormone surge and its relationship with the circadian cortisol rhythm in the human. Neuroendocrinology. 2002;75:158–163. 368. Cahill DJ, Wardle PG, Harlow CR, Hull MG. Onset of the preovulatory luteinizing hormone surge: diurnal timing and critical follicular prerequisites. Fertil Steril. 1998;70:56–59. 369. Wilcox AJ, Weinberg CR, Baird DD. Timing of sexual intercourse in relation to ovulation. effects on the probability of conception, survival of the pregnancy, and sex of the baby. N Engl J Med. 1995;333:1517–1521. 370. Rossmanith WG, Yen SS. Sleep-­associated decrease in luteinizing hormone pulse frequency during the early follicular phase of the menstrual cycle: evidence for an opioidergic mechanism. J Clin Endocrinol Metab. 1987;65:715–718. 371. Soules MR, Steiner RA, Cohen NL, Bremner WJ, Clifton DK. Nocturnal slowing of pulsatile luteinizing hormone secretion in women during the follicular phase of the menstrual cycle. J Clin Endocrinol Metab. 1985;61:43–49. 372. Mortola JF, Laughlin GA, Yen SS. A circadian rhythm of serum follicle-­stimulating hormone in women. J Clin Endocrinol Metab. 1992;75:861–864. 373. Kapen S, Boyar R, Hellman L, Weitzman ED. The relationship of luteinizing hormone secretion to sleep in women during the early follicular phase: effects of sleep reversal and a prolonged three-­hour sleep-­wake schedule. J Clin Endocrinol Metab. 1976;42:1031–1040. 374. Rahman SA, Grant LK, Gooley JJ, Rajaratnam SMW, Czeisler CA, Lockley SW. Endogenous circadian regulation of female reproductive hormones. J Clin Endocrinol Metab. 2019;104:6049–6059. 375. Klingman KM, Marsh EE, Klerman EB, Anderson EJ, Hall JE. Absence of circadian rhythms of gonadotropin secretion in women. J Clin Endocrinol Metab. 2011;96:1456–1461. 376. Lavoie HB, Marsh EE, Hall JE. Absence of apparent circadian rhythms of gonadotropins and free alpha-­subunit in postmenopausal women: evidence for distinct regulation relative to other hormonal rhythms. J Biol Rhythms. 2006;21:58–67. 377. Boyar R, Finkelstein J, Roffwarg H, Kapen S, Weitzman E, Hellman L. Synchronization of augmented luteinizing hormone secretion with sleep during puberty. N Engl J Med. 1972;287:582–586. 378. Kapen S, Boyar RM, Finkelstein JW, Hellman L, Weitzman ED. Effect of sleep-­wake cycle reversal on luteinizing hormone secretory pattern in puberty. J Clin Endocrinol Metab. 1974;39:293–299. 379. Shaw ND, Butler JP, McKinney SM, Nelson SA, Ellenbogen JM, Hall JE. Insights into puberty: the relationship between sleep stages and pulsatile LH secretion. J Clin Endocrinol Metab. 2012;97:E2055–E2062. 380. Shaw ND, Butler JP, Nemati S, et al. Accumulated deep sleep is a powerful predictor of LH pulse onset in pubertal children. J Clin Endocrinol Metab. 2015;100:1062–1070. 381. Rossmanith WG, Lauritzen C. The luteinizing hormone pulsatile secretion: diurnal excursions in normally cycling and postmenopausal women. J Gynaecol Endocrinol. 1991;5:249–265. 382. Loucks AB, Heath EM. Dietary restriction reduces luteinizing hormone (LH) pulse frequency during waking hours and increases LH pulse amplitude during sleep in young menstruating women. J Clin Endocrinol Metab. 1994;78:910–915. 383. Hall JE, Sullivan JP, Richardson GS. Brief wake episodes modulate sleep-­inhibited luteinizing hormone secretion in the early follicular phase. J Clin Endocrinol Metab. 2005;90:2050–2055. 384. Lu C, Hutchens EG, Farhy LS, Bonner HG, Suratt PM, McCartney CR. Influence of sleep stage on LH Pulse initiation in the normal late follicular phase and in polycystic ovary syndrome.

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22.e10 PART I  The Fundamentals of Reproduction 385. Collins JS, Marshall JC, McCartney CR. Differential sleep-­wake sensitivity of gonadotropin-­ releasing hormone secretion to progesterone inhibition in early pubertal girls. Neuroendocrinology. 2012;96:222–227. 386. Kim SH, Lundgren JA, Bhabhra R, et al. Progesterone-­ mediated inhibition of the GnRH pulse generator: differential sensitivity as a function of sleep status. J Clin Endocrinol Metab. 2018;103:1112–1121. 387. Whitten WK. Modification of the oestrous cycle of the mouse by external stimuli associated with the male; changes in the oestrous cycle determined by vaginal smears. J Endocrinol. 1958;17:307–313.

388. Okamura H, Murata K, Sakamoto K, et al. Male effect pheromone tickles the gonadotrophin-­ releasing hormone pulse generator. J Neuroendocrinol. 2010;22:825–832. 389. McClintock MK. Menstrual synchrony and suppression. Nature. 1971;229:244–245. 390. Stern K, McClintock MK. Regulation of ovulation by human pheromones. Nature. 1998;392:177–179. 391. Schank JC. Menstrual-­cycle synchrony: problems and new directions for research. J Comp Psychol. 2001;115:3–15. 392. Trotier D. Vomeronasal organ and human pheromones. Eur Ann Otorhinolaryngol Head Neck Dis. 2011;128:184–190.

2

The Gonadotropin Hormones and Their

2

Receptors1

Prema Narayan, Alfredo Ulloa-­Aguirre, and James A. Dias

OUTLINE INTRODUCTION GONADOTROPINS Gonadotropin Proteins (LH, CG, and FSH) Gonadotropin Structure-­Function Studies Gonadotropin Genes and Transcripts Gonadotropin Expression and Secretion Gonadotropin Clinical Significance GONADOTROPIN RECEPTORS Gonadotropin Receptor Proteins Gonadotropin Receptor Gene Expression and Regulation Gonadotropin Receptor Signaling Pathways Gonadotropin Receptor Clinical Significance Homologous Receptors Low Molecular Weight Gonadotropin Receptor Agonists and Antagonists

The three gonadotropins act via two G protein-­coupled receptors (GPCRs). The LH receptor recognizes both LH and CG (thus it is referred to as LHCGR) and the FSH receptor (FSHR) is specific for FSH. Both receptors are expressed in the male and female gonads, the LHCGR is expressed in Leydig, theca, granulosa, and luteal cells, whereas the FSHR is expressed in granulosa and Sertoli cells. Although the presence of the gonadotropin receptors in extragonadal tissues has been reported, their physiological significance is still under debate.

GONADOTROPINS • Gonadotropins are heterodimeric glycoproteins of the cystine knot family. • Their biosynthesis and secretion as well as their biological activity are dependent on glycosylation. • Their target organs are testis and ovary; they are essential for steroidogenesis and gametogenesis.

Gonadotropin Proteins (LH, CG, and FSH) Physiological Function INTRODUCTION Derived from the Greek meaning of “that which generates,” the gonads are the female and male organs that produce egg and sperm, respectively. Pituitary glycoprotein hormones which bind to target receptors on granulosa and theca cells in the ovary or Sertoli and Leydig cells in the testis have therefore been named gonadotropin hormones. The IUPAC (International Union of Pure and Applied Chemistry) nomenclature for human pituitary gonadotropin hormones are follitropin for follicle-­stimulating hormone (FSH), lutropin for luteinizing hormone (LH), and choriogonadotropin (CG) for the placental chorionic hormone. Here we will abbreviate them as LH, FSH, and CG. Unlike LH and FSH, which are expressed in mammalian and nonmammalian species, CG is expressed only in primates and equids. As implied by their names, the role of gonadotropins is to activate gonadal cells to produce oocytes and sperm necessary for procreation. In addition, they are essential for the production of steroid hormones by their target cells in gonads. Some of these steroids are essential for producing high-­quality gametes and secondary sexual characteristics associated with sexual maturity, while still others are essential for the receptivity of the uterus for implantation and maintenance of pregnancy. A unique feature of gonadotropins regardless of origin is that, at the level of primary sequence, they share one identical or common subunit encoded by a unique gene. Each one has a second unique subunit which combines with the identical/common subunit to form active trophic hormones. The common subunit has been named the alpha subunit and the unique subunit has been named the beta subunit. They are therefore heterodimeric, and incidentally, the three gonadotropins share this common alpha subunit with another pituitary hormone, thyrotropin, or thyroid-­ stimulating hormone (TSH).

The gonadotropins, LH, FSH, and human (h)CG play a critical role in the fundamental processes of development and reproduction.1,2 LH and FSH are secreted by the gonadotrope cells of the pituitary gland, whereas hCG is a placental hormone. The effects of LH and hCG are mediated by their common receptor, LHCGR, while those of FSH are mediated by its receptor, FSHR. The major function of LH in the male is to stimulate LHCGR present specifically in testicular Leydig cells to produce testosterone that is essential for the development of puberty, male secondary sexual characteristics, and spermatogenesis.3 FSH targets its receptor present in Sertoli cells of the seminiferous tubule and supports their growth and differentiation, thereby indirectly supporting spermatogenesis.4 In the ovary, LHCGR is present in theca cells lining the follicles, mural granulosa cells of the preovulatory follicle, stromal cells, and luteinized cells. Consequently, LH regulates several functions within the ovary. LH-­mediated activation of LHCGR in theca cells stimulates androgen production, while receptor activation in the mural preovulatory granulosa cells triggers autocrine and paracrine signaling pathways that lead to ovulation.5 Following ovulation, LH maintains progesterone production by the corpus luteum. FSH activates FSHR present exclusively in the granulosa cells of the follicle to stimulate the growth and maturation of the follicle, stimulate the production of aromatase for conversion of theca cell-­produced androgens to estrogen, and induce LHCGR receptors in the mural granulosa cells of the preovulatory follicle.6,7 In contrast to LH and FSH, hCG is essential for the initiation and maintenance of pregnancy and during fetal development. One of the major functions of placental hCG is to maintain progesterone production by the corpus luteum for the first few weeks of pregnancy before the transition to placental progesterone production. It also mediates multiple placental, uterine, and fetal functions including trophoblast invasion, development of syncytiotrophoblast cells, angiogenesis in

1 The authors would like to recognize the past contributions of Drs. David Puett and Mario Ascoli, who were authors of previous versions of this chapter in earlier editions of this book.

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PART I  The Fundamentals of Reproduction

hFSHβ 1 - - - - - - N S C E L T N I T I A I E K E E C R F C I S I N T TWC A GY C Y T R D L V Y K D P A R P K I Q K T C T F K E L V Y E T V R V P V C A H 68 hLHβ 1 S R E P L R PWC H P I N A T L A V E K E GC P V C I T V N T T I C A GY C P T MMR V L Q A V L P P L P Q V V C T Y R D V R F E S I R L P GC P R 74 hCGβ 1 S K E P L R P R C R P I N A T L A V E K E GC P V C I T V N T T I C A GY C P T M T R V L QG V L P A L P Q V V C N Y R D V R F E S I R L P GC P R 74 hFSHβ 69 H A D S L Y T Y P V A T QC H CG K C D S D S T D C T V RG L G P S Y C S F G EMK E - - - - - - - - - - - - - - - EMK E - - - - - - - - - 111 hLHβ 75 G V D P V V S F P V A L S C R CG P C R R S T S D CGG P K D H P L T C D H P - - - - - - - - - - - - - - - - - - - Q L S G - - - - - L L F L 121 hCGβ 75 G V N P V V S Y A V A L S CQC A L C R R S T T D CGG P K D H P L T C D D P R F QD S S S S K A P P P S L P S P S R L P G P S D T P I L P Q 145

α 1

APDVQDCPECTLQENPFFSQPGAPILQCMGCCFSRAYPTPLRSKKTMLVQKNVTSESTCCVAKSYNRVTVMG 74

75 KVENHTACHCSTCYYHKS

92

Fig. 2.1 Amino acid sequences of human α gonadotropin subunit (α-­subunit), luteinizing hormone (LH)β, chorionic gonadotropin (CG)β, and follicle-­stimulating hormone (FSH)β.  Amino acid sequences were obtained from the Ensembl website (http://www.ensembl.org/index.html), and the β-­subunits are aligned to maximize homology. Identical, highly conserved, and semiconserved residues among the three β-­subunits are highlighted by the blue, green, and yellow boxes, respectively. All cysteines participate in disulfide bond formation in the native proteins. hCG, human chorionic gonadotropin; hFSH, human follicle-­stimulating hormone; hLH, human luteinizing hormone. (Copyright 1999-­2008 The European Bioinformatics Institute and Genome Research Limited, and others. All rights reserved.)

the uterine endometrium, uterine growth and differentiation, placental development, and localized suppression of the immune system.8,9 Another important physiological function of placental hCG is in male sexual development. hCG activates LHCGR in the fetal Leydig cells to produce testosterone that stimulates the growth and differentiation of male genitalia.10 The fetal ovary is not sensitive to hCG and female sexual differentiation is independent of gonadotropins.

C C

N

Protein Structural Attributes The three gonadotropins and TSH comprise the better characterized members of a family of complex proteins known as the glycoprotein hormones.1 They are noncovalently bound heterodimers composed of a common α-­subunit and distinct β-­subunits. The common α gonadotropin subunit (α-­subunit) contains 92 amino acid residues, and LHβ, FSHβ, and hCGβ subunits are, respectively, 121, 110, and 145 amino acid residues in length. The additional length of the hCGβ subunit is due to a carboxy-­terminal extension arising from a frameshift mutation in an ancestral LH β-­subunit gene resulting in a read-­through into an untranslated region of the LHβ subunit and an extension of the open reading frame.11–14 This extension is known as the carboxy-­terminal peptide (CTP). The amino acid sequences of the human subunits are shown in Fig. 2.1, and it can be seen that the α and β-­subunits are relatively rich in Cys residues and that considerable homology exists in the β-­subunits. Crystal structures have been determined for partially active deglycosylated hCG,15,16 glycosylated, antibody-­bound hCG,17 a partially deglycosylated fully active hFSH,18 and a partially deglycosylated complex of a single chain hFSH bound to a large N-­terminal fragment (residues 1-­268) of the hFSHR ectodomain (ECD)19 and FSH-­FSHR complex containing the entire ECD including the hinge region that is required for signal specificity.20 Fig. 2.2 shows the crystal structures of hCG and hFSH. Recently, the crystal structure of bovine (b) LH β was reported.21 The conformations of hCG and hFSH are quite similar, each being highly elongated molecules with the two subunits intertwined one with another in a slightly twisted manner. Despite the absence of any striking sequence homology, the two subunits in both heterodimers have similar folds characterized by three major loops, and each subunit contains a cystine knot motif, consisting of three disulfides located in the core of each subunit. The α and β-­subunits contain, in addition to the three disulfides in the cystine knot, two and three disulfides, respectively. A 20-amino acid residue region of the

N

N

C

N

C

Fig. 2.2 Crystal structures of human chorionic gonadotropin (hCG; left) and human follicle-­stimulating hormone (hFSH; right). The structures15–18 show that the two subunits are highly elongated and intertwined (α-­subunit, yellow; hCGβ, green; and FSHβ, blue) forming a relatively large contact surface area. As discussed in the text, there are several interesting features associated with the structures: each subunit contains a cystine knot motif; the β subunit wraps around a portion of the α subunit forming what is termed a seatbelt (shown in white); although having little sequence homology, the two subunits adopt similar folding patterns; hCG and hFSH form very similar structures, but the respective subunits of each exhibit subtle differences in their conformations. C, C-­terminus; N, N-­terminus.

around a portion of the α-­subunit like a molecular seatbelt held in place with disulfide bonds. A major difference in the structures of hCG and hFSH is in the C-­terminal portions of the seatbelts that exhibit distinct conformations. In both hCG and hFSH, the two subunits are associated in a head-­to-­tail arrangement (Fig 2.2). While the structures of bLHβ and hCGβ are similar in the cysteine-­knot core, the conformations of the extended loops show variation.21 Solution structures have also been obtained for deglycosylated human α-­subunit22,23 using NMR spectroscopy. The overall ensemble of structures determined for the α-­subunit is similar to that obtained in the crystal structures of hCG and hFSH.

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

α-Subunit H2N (92 aa) SO4 SO4

N 30

LHβ H2N (121 aa)

FSHβ (110 aa)

CGβ H2N (145 aa)

N 52

H2N

N 7

N 13

N 24

N 30

N 78

COOH

25

N-acetyl-glucosamine Mannose Galactose Sialic acid N-acetyl-galactosamine

COOH

COOH

O O

O O COOH

138 121 127 132

Fig. 2.3 Location and typical structures of the N-­linked and O-­linked glycans on the gonadotropins. The sites of glycosylation on each of the gonadotropin subunits and representative structures of the various N-­linked glycans on human luteinizing hormone (hLH), human chorionic gonadotropin (hCG), and human follicle-­stimulating hormone (hFSH). As discussed in the text, scores of different structures have been identified, often with fucose present; moreover, some sites are devoid of glycosylation in various gonadotropin preparations. A representative structure of type 1 O-­linked glycans on the carboxy-­terminal peptide of hCGβ is shown; type 2 structures have been reported as well, and some sites show no glycosylation in some gonadotropin preparations. Blue squares, GlcNAc; yellow squares, GalNAc; green circles, mannose; yellow circles, galactose; purple diamonds, sialic acid.

With crystal structures available for FSH18 and the FSH-­ FSHR ECD complex19,20 (discussed elsewhere in this chapter), it is possible to delineate the conformational changes of the free heterodimer and that bound to a receptor. The unbound form of the hormone is more flexible than that of the bound form. It is the C-­terminal region of the α-­subunit; however, that undergoes the greatest change in conformation. In addition, the two C-­terminal residues in the α-­subunit are unordered in the crystal structure of FSH, but they are fully ordered in the complex with receptors.

Glycosylation During their synthesis, gonadotropins are trafficked from the endoplasmic reticulum to the cis-­Golgi and undergo glycosylation as they traverse the Golgi reaching the trans-­Golgi, to yield the mature hormones. The human subunit primary sequences contain N-­linked glycosylation sites (consensus sequence Asn-­ X-­Ser/Thr, where X is any amino acid except proline): two on α-­subunit at Asn52 and Asn78, two on hCG β-­subunit (Asn13 and Asn30), two on FSHβ (Asn7 and Asn24) and one on LH β-­ subunit at Asn30. In addition, the hCG β-­subunit contains four mucin-­type O-­linked glycans at serine 121, 127, 132, and 138, located on the CTP (Fig. 2.3), resulting in a longer half-­life of hCG as compared to LH.12 The carbohydrate moieties appear to be important in subunit assembly and stabilization, secretion, and circulatory half-­life. Although earlier studies suggested a role of the N-­linked glycan at Asn52 on

evidence indicates that, in addition, the glycan acts as a conformational or stabilizing determinant of the protein.24,25 Moreover, there is growing evidence that the particular type of glycosylation may influence biological activity.26–34 Of note is that the oligosaccharides on the α-­subunit differ in a hormone-­specific manner, apparently influenced by its cognate partner, since the characterization of the oligosaccharides released from the α-­subunit can identify the β-­subunit with which it was associated.35–37 The biantennary N-­linked glycans on hFSH and hCG terminate in sialic acid (and sulfate to a lesser extent), and the number of such moieties varies from 0 to 2, accounting in large part for the microheterogeneity of these glycoprotein hormones. In LH, the biantennary N-­linked structures tend to terminate mainly in sulfate resulting in a decrease in its circulatory half-­life compared to the sialic acid-­containing hormones. This arises from a hepatic receptor that recognizes the terminal N-­acetyl galactosamine-­sulfate, rapidly removing it from circulation.38 Indeed, ablation of the gene encoding GalNAc-­4-­sulfotransferase, the enzyme responsible for modifying the terminal GalNAc in LH with sulfate, resulted in mice with increased half-­life and circulating levels of LH.39 These are but generalizations since, for example, hFSH also contains triantennary and tetraantennary N-­linked glycans, and some hFSHβ-­subunits lack N-­linked structures completely.40–44 A variety of glycosyltransferases are responsible for N-­and O-­glycan biosynthesis; notably, sulfation in the pituitary requires N-acetylgalactosamine transferase and sulfotransferase, both of Also, fucose is often found

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PART I  The Fundamentals of Reproduction

in the glycoprotein hormones. As summarized, nearly 50 different N-­linked and O-­linked glycans have been reported in hCG α and β preparations.45–47 The major gonadotropin N-­linked and C-­linked glycans are shown in Fig. 2.3.

Folding and Assembly Investigations into the kinetic folding pathways of the hCG subunits led to the interesting suggestion that disulfide exchange occurs during the maturation process and that subunit association occurred before completion of protein folding and disulfide formation.48–52 Moreover, it was posited that subunit association occurred before the seatbelt was latched by closure of Cys26 and Cys110 (wraparound model). In contrast to these reports, a different mechanism in which subunit assembly involved closure of the seatbelt latch, followed by threading of α loop 2, has been proposed (threading model).53–57 Others have also studied the folding patterns of hCG and reported that subunit association occurred between an almost completely folded α-­subunit and an immature β-­subunit.58–61 Another study suggests that the LH β-­ subunit is not completely folded prior to assembly with the α-­ subunit and that the α-­subunit serves as a chaperone to facilitate the formation of the cysteine knot and the seatbelt latch.62 In addition to the heterodimeric nature of the hormones, homodimers have also been found for the LH β-­subunit63,64 and for the α-­subunit.58,59,65 Whether these homodimeric forms of the glycoprotein hormones have any associated bioactivity remains to be shown, although it has been reported that the free α-­subunit potentiates progesterone-­mediated decidualization.66

Gonadotropin Structure-­Function Studies • Prior to structural determination, extensive structure-­activity relationships were established through mutagenesis, which ultimately confirmed the authenticity of the gonadotropin-­ receptor crystal structures. • In addition, these mutants established the essentiality of posttranslational modifications on the assembly, secretion, and biological activity of the gonadotropins. • Protein engineering efforts have been directed at producing longer-­ acting and superagonist forms of gonadotropins.

Site-­Directed Mutagenesis As discussed elsewhere in this chapter, only a limited number of naturally occurring mutations have been identified in α and β-­subunits of the glycoprotein hormones. In contrast, there is a wealth of information available from site-­directed mutagenesis followed by biological characterization of the mutant hormones. Few mutants have been described in which there was a significant increase in bioactivity; most mutations either have no effect or induce a loss of function, either disrupting folding, subunit assembly, or receptor binding/activation. Gain-­of-­function mutations in hCG were obtained by replacing single or multiple amino acid residues at the N-­terminal region of the α-­subunit with Lys.67 A twofold increase in potency of hCG was obtained with a single replacement of Phe with Thr at position 18 of α-­ subunit.68 Mutant forms of α-­subunit missing the N-­linked oligosaccharide at Asn52 are capable of associating with the hCG β-­subunit or FSHβ-­subunit giving a heterodimer that binds to the cognate receptor but has diminished signaling efficacy.69 It has been suggested that the role of N-­linked glycosylation at Asn52 is to stabilize the active conformation of the heterodimer by formation of a hydrogen bond with a Tyr on the β-­subunit.18 Mutations in the central region70–72 and at the C-­terminus73,74 of α-­subunit yielded mutants that associated with the β-­subunits of hCG and hFSH but displayed compromised functionality in receptor binding.

A large number of β-­subunit mutations have been prepared and characterized.75–82 As with mutations in α, many interfere with folding, subunit assembly, or receptor binding. Overall, the data is consistent with the crystal structures of hCG and FSH. Deletion mutants at the N-­and C-­termini of the hCG β-­subunit have also been reported by several groups75,77,79,83 and the shortest form that retains minimal functionality in subunit assembly and subsequent receptor binding and activation is a fragment consisting of residues 8–100.77 Unlike α-­subunit mutants, however, there have been no reports of β-­subunit mutants that retain the ability to form heterodimers and bind to receptors but do not signal. This suggests that the α-­subunit may play a predominant role in the activation of gonadotropin receptors following the initial binding event.

Protein Engineering Protein engineering has been used to produce a variety of chimeric and single-­chain hormones yielding quite interesting results. A number of glycoprotein hormone chimeras have been characterized, providing useful information on specific amino acid residues involved in receptor binding and activation.84–90 In general, these results emphasize the role of the β-­subunit seatbelt region in receptor binding, although different portions are important in receptor specificity. A novel approach to studying gonadotropin structure-­ function relationships involved the design of single-­chain (yoked or tethered) hormones, derived by fusion of α and β-­subunits. In the configurations N-­hCGβ-­α-­C or N-­α-­hCGβ-­C, single-­chain hCG with or without intervening peptide linkers were expressed and characterized, in many cases with interesting mutations in one or both subunits.91–95 From these studies, it was concluded that both subunit configurations were bioactive and, quite surprisingly, that, each disulfide of the subunits could be eliminated without a loss of activity.96,97 Subsequently, similar fusion proteins of LH and FSH were also found to be bioactive.98,99 The single-­ chain hormones displayed increased stability and heat resistance in vitro compared to their heterodimeric counterparts.98 Extending the approach of covalently linking the two subunits, disulfide-­linked heterodimers and mini-­gonadotropins with full bioactivity were designed and expressed.25,100–104 These gonadotropin analogs substantiated the notion that α-­Asn52 contributed to heterodimer stability and was not involved directly in signal transduction25 and led to suggestions that the C-­terminus of the α-­subunit is not required for LHR binding. This notion was dispelled by observations that some but not all single amino acid mutations in the C-­terminus of the α—subunit in the context of heterodimeric hFSH only retained 10% or less of hFSH receptor-­binding activity.105 The use of single-­chain gonadotropins, particularly in the N-­α-­β-­C configuration, also raises interesting questions about the role of the C-­terminal region of the α-­subunit in FSH where the structure of the hFSH-­hFSHR ECD complex shows a large movement of α-­subunit in the receptor complex compared to the heterodimer.19 This riddle is unlikely to be solved until the crystal structure of single-­chain gonadotropin in complex with receptor is determined. Of great interest was the report that single-­chain hCG β-­β homodimers bind to LHCGR with an affinity about three times lower than wild type hCG but do not elicit a biological response and block hCG binding to LHCGR.106 A second single-­chain hCG antagonist was designed by mutating three of the four N-­linked glycosylation sites that are associated with LHCGR activation (Asn 13 and 30 in the β-­subunit and Asn 52 in the α-­subunit). This analog behaved as a competitive antagonist and suppressed ovarian hyperstimulation syndrome in rats.107 The single-­chain methodology has been extended to produce fusion proteins with dual and triple activities, although their mechdomain fusion

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

protein of the form, N-­FSHβ-­hCG β-­subunit-­α-­C, exhibited both LH and FSH activities.108–111 Disruption of heterodimer formation in this triple domain construct by mutation of either Cys10-­ Cys60 or Cys32-­Cys84 did not eliminate bioactivity, suggesting that αβ contacts are not required for receptor binding and activation.112 Subsequently, a four-­domain fusion protein, N-­TSHβ-­ FSHβ-­hCG β-­subunit-­α-­C, although secreted inefficiently, was found to exhibit three distinct bioactivities both in cellular and whole animal studies.113,114 These results raise intriguing questions regarding subunit association and conformation as manifested in receptor binding and activation. These selected results, along with others not covered here, indicate that single-­chain glycoprotein hormones exhibit some properties distinct from those in heterodimers. The increased stability of the single-­chain proteins, the single-­chain hCG antagonists, and the analogs with dual activities make them interesting candidates for clinical utility. The approach of single-­ chain gonadotropins was further expanded to produce fusion proteins of single-­chain hCG with LHCGR, which when expressed, led to constitutive receptor activation in transfected cells and transgenic mice.115–119 This model also demonstrated that protein fusions of the individual subunits with LHCGR were devoid of bioactivity.120

Gonadotropin Genes and Transcripts • Except for the hCG β-­ subunit, the gonadotropin subunits are encoded by one gene. Not unexpectedly, mutations in gonadotropin genes discovered in symptomatic patients often affect assembly, conformation, and thus biological activity. • Polymorphisms in the protein coding regions of gonadotropin genes can similarly have detrimental or no effects depending on whether the polymorphism results in an amino acid change or is silent, and whether the change in primary structure affects function. • In addition, polymorphisms in noncoding regions can affect transcription or mRNA processing, which may or may not be associated with clinical conditions.

Gonadotropin Subunit Genes The common α-­subunit and the β-­subunits of LH and FSH are each encoded by single genes; in contrast, the CG β-­subunit, expressed in primates, is encoded by six genes.121,122 In equids, however, the CG β-­subunit and LH β-­subunit are products of the same gene.123 It has been suggested that the glycoprotein hormone α and β-­subunits diverged from a common ancestral gene over 900 million years ago,121 with the β gene undergoing duplications and mutations to yield the current family. In humans, the gene encoding the common α subunit, CGA, is on chromosome 6, which for FSHβFSHB) on chromosome 11 and those for LHβ (LHB) and CGβ(CGB) on chromosome 19 (http://www. ensembl.org). The CGA is 9.4 kb and contains four exons and three introns, FSHB is 4.2 kb with three exons and two introns, LHB is 1.1 kb with three exons and two introns, and the CGB genes are variable in length. The one LHB and six CGB genes exist in a large cluster spanning about 52 kbp.121,122 The six CGB genes (i.e., CGB, CGB1, CGB2, CGB5, CGB7, and CGB8) exist as tandem and inverted repeats.124 Detailed analysis of the CGB genes revealed that four of the genes, CGB, CGB5, CGB7, and CGB8, exhibit 97% to 99% sequence identity while their identity with LHB gene is 92% to 93%.121,122 These gene similarities lead to protein sequences that are 98% to 100% identical for the four CG β-­subunits and 85% identical with the LH β-­subunit.

Gonadotropin Subunit Transcripts The available evidence indicates single transcripts for the gonad otropin genes, with the exception of the human

27

which four mRNA species have been described, arising from alternate splicing and the utilization of two polyadenylation sites.125 The CGB family is interesting in that the six genes appear to express transcripts of varying lengths, albeit sometimes without detectable protein production. CGB5 and CGB8 are highly expressed in the placenta.121,122,126 CGB1 and CGB2 genes are expressed in placenta,121,122 pituitary,127 testis,128 and in breast cancer,129 although no proteins have yet been identified for these genes. The predicted sizes of putative protein products of CGB1 and CGB2 are smaller than that of the hCG β-­subunit; this observation, coupled with the distinct amino acid sequences predicted, suggests that, if biosynthesized, these proteins may have quite different functions than those of hCG. Using transgenic mice expressing a 36 kb cosmid insert that contained the six CGB genes, transcripts of CGB1 and CGB2 genes were found to be present in the brain at levels comparable to those of the other four CGB genes.130 The human LHB mRNA is 700 nucleotides in length, and depending upon the species, the CGA gene encodes an mRNA of 730 to 800 nucleotides.125

Naturally Occurring Mutations Mutations in the gonadotropin hormone genes, although rare, help in elucidating their physiological roles and defining the structural domains of the hormones. The only mutation reported in the CGA gene is that from a human carcinoma, Glu56Ala, resulting in a mutant form of α-­subunit that does not associate with LH β-­subunit.131 In contrast, there are several reports of mutations in the genes encoding the three gonadotropin β-­subunits, resulting in loss of function and thus hypogonadism.122,132,133 The first report of a mutation in the LHB gene was that of a missense mutation in a male presenting with delayed puberty and hypogonadism.134 This mutant led to a replacement of Gln54 with Arg; while subunit assembly could occur, the heterodimer was unable to bind to LHCGR. Site-­directed mutagenesis studies showed that LH β-­subunit and hCG β-­subunit with Gln54 replacements formed heterodimers with α-­subunit, but these heterodimers exhibited reduced binding to LHCGR.76,78 Another missense mutation reported in LH β-­subunit was that of Gly36 to Asp, reported in a male with delayed puberty and infertility.135 Gly36 is part of the CAGYC sequence in the LH β-­subunit that is critical to the formation of the cystine knot; presumably, an Asp at this position prevents at least one of the disulfides from forming. The third identified mutation was a G-­C substitution at the +1 position of intron 2 (a 5′ splice-­donor site) that leads to a hypothetical aberrant protein with a 79 amino acid residue insert beginning after Met41 and a frameshift in exon 3, thus removing the essential seat belt loop of β and cysteines that participate in the cysteine knot motif.136 This suggested that the mutant LH-­ β subunit would not correctly assemble with the α-­subunit and therefore would not be secreted. The offspring of consanguineous parents who were heterozygous for the mutation were analyzed. Three homozygous siblings presented with hypogonadism and infertility, undetectable levels of LH, and high levels of free α-­subunit while their heterozygous siblings were fertile. The two homozygous males had elevated FSH and low testosterone. The homozygous female had FSH, estradiol, and progesterone values in the normal range and underwent normal pubertal development and menarche at age 13 years followed by oligomenorrhea and anovulation. A 9 bp deletion in exon 2 resulting in deletion of amino acid residues 10 to 12 of LHβ was reported in a man and his sister.137 Both were homozygous for the deletion, while two additional unaffected siblings were heterozygous. In spite of undetectable levels of LH and concomitant low serum and intratesticular testosterone concentrations, the man had complete spermatogenesis and normal sperm count. Presumably, the low activity by the mutant LH

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PART I  The Fundamentals of Reproduction

detected in vitro was sufficient for normal spermatogenesis. The sister underwent normal puberty and menarche but subsequently had amenorrhea, infertility, ovarian cysts, and low estradiol levels. A compound heterozygous mutation in a 31-­year-­old male with delayed puberty, azoospermia, and hypogonadism due to lack of LH was reported.138 The first mutation was identified as a 12-­bp deletion in exon 2 of the LHB gene, causing a deletion of 4 leucine residues in the signal peptide and the second was a G to T mutation at the 5′ splice site of intron 2 resulting in aberrant RNA splicing. The patient’s 16-­year-­old sister harboring the same mutations had normal pubertal development but developed oligomenorrhea. A homozygous 3 bp deletion in the LHB gene resulting in the deletion of Lys40 in LHβ was identified in two brothers with LH deficiency and hypogonadism. The mutated LHβ-­subunit was able to heterodimerize with α-­subunit but was not secreted.139 Two cases of male hypogonadism caused by premature termination of LHβ have been reported.140,141 Homozygous deletion of a thymine nucleotide at position 325 in exon 3, predicted to result in frameshift and premature termination at codon 128 was identified in a 19-­year-­old man with delayed puberty140 and a homozygous mutation of c.84G>A[p.W28X] produces a truncated protein of seven amino acids.141 Rare missense heterozygous mutations in the hCGβ subunit have been identified in a Northern European population.122,142 A Val56Leu mutation in the CGB5 gene was identified in a patient with recurrent miscarriage (RM). Although the mutation impaired subunit assembly, it elicited a strong signaling response.142 An amino-­terminal Arg8Trp mutation, also found in an RM patient, did not affect assembly. A Pro73Arg mutation found in 5 individuals (3 RM and 2 controls) resulted in altered conformation but did not affect biological activity. Individuals homozygous for these mutations have not been identified, perhaps because such genotypes would result in complete pregnancy failure and suggests that only mutations with mild functional consequences can be tolerated in the major CGB genes. To date, 13 patients with mutations in exon 3 of FSHB have been identified and they present with an absence of pubertal development, amenorrhea, and infertility in females and delayed or normal puberty with azoospermia and infertility in males.122,132,133 The first reported mutation was a homozygous 2 bp deletion at codon 61 (Val61X) causing a frameshift and premature termination of the β subunit in two women and later also identified in an 18-­year-­old male with delayed puberty.143–145 These observations were followed by a report of a compound heterozygous mutation, with one being the Val61X mutation and the other a missense mutation resulting in a Cys51Gly replacement.146 Other identified mutations include nonsense mutations (Tyr76X and R115X),146–149 a frameshift mutation at codon 79 (Ala79X) caused by a 1 bp deletion resulting in premature termination,150 and two missense mutations resulting in Cys82Arg and Cys122Arg replacements.151,152 These mutations result in loss of bioactivity due to the production of a truncated protein or due to aberrant tertiary structure as a result of mutations in the cysteine residues involved in the cystine knot structure and inability to associate with the α-­subunit. One case of hypoglycosylation, likely caused by altered conformation, was reported for FSH, resulting in a hormone with diminished activity.153 Overall, the observed phenotypes associated with the naturally occurring mutations in LHB, FSHB, and CGB are consistent with the known structures and actions of the gonadotropins, although fertility in men does not always appear to be sensitive to some FSH mutations.

Polymorphisms A well-­characterized variant in the LHB gene (V-LHβ) appears in variable frequencies in ethnic groups throughout the world

and results from two single nucleotide polymorphisms (SNPs) that are found together on one allele. One causes the replacement of Trp8 with Arg which resulted in altered immunoreactivity of the hormone. The other caused a substitution of Ile15 with Thr, which introduces an extra glycosylation site in the LHβ subunit.122,132,133 V-­LH demonstrates increased biopotency in vitro with an altered half-­life in circulation.154,155 The association of the V-­LH with various clinical conditions has been asses sed.122,132,133 A number of studies have addressed the association between V-­LH and various clinical conditions such as infertility, polycystic ovarian syndrome, and menstrual disorders. No clear association was found with PCOS.122 However, studies have found an association with female infertility, but not male.156–159 Another LH β-­subunit variant with a replacement of Gly102 with Ser and resulting in reduced LH biopotency in vitro has been associated with reproductive disorders in some populations.122,132,133 The frequency of this polymorphism was recently reported to be higher in a population of Chinese Han women with PCOS but not in a population of Korean women.160,161 An unusual polymorphic variant of LH β-­subunit involves an Ala to Thr replacement of three residues before the signal peptide cleavage site.132,133 Using in vitro assays, it was found, rather surprisingly, that the mature protein from the variant appears less potent than wild type LH in cAMP production but more potent in inositol phosphate production. The SNP-­related alteration may interfere with the proper processing of the β-­subunit, although studies have not addressed this possibility. A polymorphism in exon 3 of CGB5, resulting in a Val79 replacement with Met in a random population in the United States, has been reported.162 This SNP results in a β-­subunit with impaired ability to assemble with the α-­subunit, although the biological activity of the variant is normal. The frequency and physiological consequences of this polymorphic variant are unknown; one sampling of just under 600 samples from four European groups failed to detect the polymorphism in this population.163 Other polymorphic variants have been detected, but these are silent or located in intron regions.132,133 A case study analyzed CGB5 and CGB8 genes in RM and control fertile patients from Estonia and Finland. Seventy-­one polymorphisms were identified, of which 48 were novel.164 A protective effect against RM was associated with two SNPs located at identical positions in CGB5 and CGB8 and with four CGB5 promoter variants. A follow-­up study that included a Danish cohort with RM in addition to the Estonian and Finnish subjects confirmed that two SNPs in the CGB5 promoter region seemed to offer protection against RM, but variants in the CGB8 promoter region had no effect.165 These polymorphisms can be found in the dbSNP database (http://www.ncbi.nlm.nih.gov/SNP/). Only a few SNPs in the FSHB gene have been extensively studied.122,166 An FSHB promoter polymorphism (rs10835638) (G/T) located 211 bp upstream of the transcription start site was identified in a cohort of European men and TT homozygous men have reduced serum FSH levels.167 This polymorphism has been associated with infertility in both sexes.168–171 The low-­ serum FSH levels associated with this SNP have been demonstrated to be due to reduced binding of the LHX3 transcription factor and reduced FSHB transcription.172 Recent genome-­wide association studies (GWAS) identified an SNP (rs11031006) (G/A) associated with fertility and PCOS. This SNP was located approximately 26 Kb upstream of the FSHB transcriptional start site.173,174 Surprisingly, functional studies revealed that the SNP resides within a conserved enhancer region and the minor (A) allele increased SF1 binding to the enhancer and increased FSHB transcription instead of decreasing expression as would be predicted in PCOS.175 The LHB and FSHB polymorphisms can be found in the SNP database (www.ncbi.nlm.nih.gov/projects/ SNP/snp_ref.cgi?geneId=3972 and www.ncbi.nlm.nih.gov/proj-

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

Gonadotropin Expression and Secretion • Production of biologically active gonadotropin is regulated in the hypothalamic-­pituitary axis at the level of transcription by trophic factor initiated intracellular signaling pathways. • Whereas regulation of transcription is by transcription factors, it is glycosylation and its link to secretion that ultimately determine the production of cognate protein. • Those processes are dependent on and guided by appropriate nascent protein primary structure.

Transcriptional Regulation The neuroendocrine reproductive axis, composed of the hypothalamus, anterior pituitary, and gonads, is regulated in large part by kisspeptin, a product of KISS1, which acts via the GPCR, KISS1R, to stimulate the gonadotropin-­ releasing hormone (GnRH) neurons in the hypothalamus.176,177 The three gonadotropin subunit genes in the pituitary gonadotropes are differentially responsive to GnRH pulse frequency and magnitude: LHB is preferentially transcribed at high GnRH pulse frequencies and FSHB at lower frequencies. The common alpha subunit is produced in excess of LHβ and FSHβ regardless of GnRH pulse frequency.178,179 Therefore, control of LH and FSH synthesis is correlated with the expression of LHβ and FSHβ subunits. Sex steroid-­mediated feedback regulation occurs mostly at the hypothalamic level, although there are also some direct actions at the pituitary and recent evidence suggests a critical role of the kisspeptin-­KISS1R system in sex steroid action.180,181 Our understanding of the signaling mechanisms that mediate expression of the pituitary gonadotropin genes has primarily come from studies on the rodent genes in two murine immortalized gonadotrope cell lines, αT3-­ 1 and LβT2, and from in vivo studies in mouse models.182,183 The binding of GnRH to its receptor (GnRHR) in the gonadotropes activates the classic Gαq/11 pathway, resulting in the stimulation of phospholipase Cβ, inositol triphosphate mediated increase in intracellular calcium from endoplasmic reticulum stores, and activation of protein kinase C (PKC) and calcium-­calmodulin kinase II. GnRHR activation also mediates calcium influx via L-­type voltage-­gated calcium channels.184,185 PKC mediates the downstream activation of the mitogen-­activated protein kinase (MAPK) cascades: extracellular signal-­regulated kinase (ERK1/2), jun N-­terminal kinase (JNK), and P38.185–187 GnRHR also couples with Gαs to activate the cAMP/PKA pathway. Therefore, GnRHR couples to different G proteins and may differentially activate distinct signaling pathways and transcriptional mechanisms in response to varying GnRH pulse frequencies for the differential synthesis of LHβ and FSHβ.188,189 Studies suggest that the ERK1/2 pathway plays a role in both Lhb and Fshb gene expression, while PKA-­mediated signaling occurs at low pulse frequencies and plays a role in Fshb expression.190–192 Higher GnRH pulse frequency appears to preferentially increase CaMKII activity, resulting in greater Lhb than Fshb expression.179 Several excellent detailed reviews on the GnRH and steroid-­ mediated transcriptional regulation of Lh and Fsh genes are available.185,188,193–196 Briefly, GnRH activation of MAPK signaling cascades increases transcription of the immediate early genes including Egr1, Jun, and Atf3 which encode the DNA proteins early growth response protein 1 (EGR1), JUN, and activating transcription factor 3 (ATF3), respectively. EGR1, SF-­1, and PITX1 form a tripartite complex that binds to a highly conserved proximal promoter sequence synergistically activating the LHB/ Lhb gene.185,188,189 Activator protein 1 (AP1), NR5A1, nuclear factor Y, PITX1, and LHX3 are some of the factors implicated in Fshb expression.185,188,194,195 β-­ catenin regulates both Lhb and Fshb in response to GnRH in vitro 197 198 although it is not essential for gonadotropin synthesis

29

forkhead box transcription factor FOXO1 inhibits Lhb and Fshb transcription in LβT2 gonadotrope cells.200,201 Activin, a member of the transforming growth factor β(TGFβ) superfamily is produced by the pituitary gonadotropes and is an important autocrine regulator of Fshb gene expression.202 Activin signaling is mediated by the activin type II receptor203 and the transcription factor FOXL2 is required for activin responsiveness.204–207 Cell culture and in vivo studies using knock-­out mice have demonstrated that SMAD 4 and FOXL2 function synergistically to regulate mouse Fshb and human FSHB transcription.208–213 Inhibin and follistatin are antagonists of activin and together the three proteins regulate FSH production at the level of transcription.188,194 Studies in the gonadotrope cell lines have demonstrated that the expression of Cga, Lhb, and Fshb are also controlled by epigenetic mechanisms that include histone modifications and alterations in chromatin structure.214,215 It has been proposed that in the trophoblast layer of the placental villous, an association of the cAMP response element binding protein (CREB) and ETS2 is augmented by protein kinase A and thereby regulates CGA expression. An upstream regulatory element on CGA contains binding sites for several transcription factors while a second control element, α-­ACT, binds a GATA factor and AP2γ.216,217 The minimal promoter of CGB lacks consensus sequences such as CAAT and TATA box. Expression of CGB is regulated by transcription factors such as AP2, SP1 and SP3, and OCT4.218–221 During pregnancy, placental CGB expression is dependent on epigenetic changes and regions of DNA in the CGB gene cluster are hypomethylated.222

Posttranslational Regulation (Glycosylation) As discussed earlier, gonadotropin carbohydrate structure is highly variable and may be the result of microheterogeneity due to structural heterogeneity of carbohydrates at the same site or macroheterogeneity due to the absence of one or more glycan chains at known glycosylation sites.42 It is now well established that carbohydrate microheterogeneity of glycoprotein hormones can vary with physiological states.28,223 Examples include a shift in the structures of hCG N-­linked oligosaccharides during pregnancy in the differentiation of cytotrophoblasts to syncytiotrophoblasts,26,224–227 changes in LH and FSH N-­ linked glycosylation during the menstrual cycle and with increasing age,42,228 as well as alterations in FSH N-­linked glycans during adolescence in boys.28,229 Oligosaccharide complexity of recombinant hFSH has been shown to differentially affect Sertoli cell endocrine activity, steroidogenesis, and gene expression in human granulosa cells.223,230 Various laboratories have shown that the hyperglycosylated variant of hCG is produced from cytotrophoblasts in early pregnancy and in gestational trophoblastic diseases, whereas hCG is secreted by the syncytiotrophoblast.224,231–233 Hyperglycosylated hCG molecules have enhanced branching into triantennary and unusual biantennary N-­linked oligosaccharides and a prevalence of the more complex Core 2, as opposed to Core 1 O-­linked sugars.232,234 In clinical studies hyperglycosylated hCG is identified by the binding of a monoclonal antibody B152 that specifically recognizes the Core 2 O-­linked sugars at Ser 132.23,236 It has been suggested that in addition to the canonical physiological function of rescuing the corpus luteum, hyperglycosylated hCG acts in an autocrine/paracrine manner to promote trophoblast invasion of the placenta.26,237–239 There are also data suggesting that the hCG produced by cytotrophoblasts and choriocarcinoma have distinct carbohydrate moieties.240 The N-­linked oligosaccharides on hCG from the invasive mole and testicular cancer are characterized by both biantennary and triantennary structures and often more heavily fucosylated glycans, while the four O-­linked units

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PART I  The Fundamentals of Reproduction

A hFSH24

B hFSH21

FSH

FSH 52

78

24

78

24

FSH 52

FSH 21 7

D hFSH15

FSH 52

FSH 24 7

C hFSH18

78

52

78

FSH 18 7

24

FSH 15 7

24

Fig. 2.4 Models of hFSH glycosylation variants and structures of the most abundant glycans. The top panel shows models of (A) hFSH24 with all four N-­glycans (B) hFSH21, which lacks the Asn24 glycan on FSHβ (C) hFSH18, which lacks the Asn7 glycan on FSHβ and (D) hFSH15, which lacks both glycans. The polypeptide backbones of α-­subunit (green) and FSHβ (blue) are shown. The N-­linked glycans are shown as spheres and represent the most abundant glycans. The bottom panel shows a symbolic depiction of the most abundant glycan at each glycosylation site. Blue squares, GlcNAc; green circles, mannose; yellow circles, galactose; purple diamonds, sialic acid; red triangles, fucose. (Figure kindly provided by Dr. George Bousfield, Wichita State University and modified from Davis JS, Kumar TR, May JV, Bousfield GR. Naturally occurring follicle-­ stimulating hormone glycosylation variants. J Glycom Lipidom. 2014;4(1):e117

Samples from patients with choriocarcinoma, testicular cancer, and invasive mole showed interesting differences in their glycans complexity. Triantennary N-­linked glycans increase in choriocarcinoma232 at Asn30 but not at Asn13,47 while monoantennary N-­linked glycans were observed at both Asn13 and Asn30.47 The status of hCG fucosylation in pregnancy and cancer patients has been investigated by several groups241 with some conflicting results. In malignancies, fucosylation was reported to increase at Asn13 but not at Asn30.47 Four naturally occurring human pituitary FSH glycosylation variants have been identified based on the loss of one or more N-­glycans in FSHβ.43,242,243 These glycoforms are named for their approximate molecular mass of the FSHβ subunit based on mobility in western blot analysis with reference to marker protein standards. A depiction of what these forms may look like is represented in Fig. 2.4. The fully glycosylated FSH possessing N-­glycans at both Asn7 and Asn24 migrates as a 24 kDa band and is designated as hFSH24 while the absence of both glycans produces a 15kDa band (FSH15). Single hypoglycosylated variants are a mixture of FSH18 and FSH21 and represent loss of glycosylation at Asn7 and Asn24, respectively. Three variants, FSH18, FSH21, and FSH24, are secreted and most pituitary and urinary hFSH preparations consist of a mixture of the more abundant hFSH24 and hFSH21 in an 80:20 ratio.43,243 Deglycosylated hFSH15 is not secreted and in vivo studies utilizing Fshb null mice have shown the N-­linked glycans on the

for efficient assembly with the α-­subunit in the pituitary and subsequent secretion.244 A progressive decrease of FSH21 has been detected in women between the ages of 24 and 55, resulting in a decrease in the FSH21 to FSH24 ratio.242 This decrease in FSH21 suggests a loss in biological activity of the mixture of circulating hFSH. Hypoglycosylated hFSH containing a mixture of FSH18 and FSH21 has higher receptor binding activity and is more potent in stimulating the cAMP pathway and steroidogenesis in a human granulosa tumor cell line and in porcine primary granulosa cells compared to FSH24.29–31,245 Recent studies suggest that FSH21/18 and FSH24 may activate distinct biological pathways in vivo, in cell culture, and in preantral ovarian follicles.30,32–34 It has been proposed that the loss of the hypoglycosylated FSH may contribute to the loss of ovarian function associated with aging.245

Regulation of Secretion Pathways of secretion and polarity of hormone release are different for the various gonadotropins. Although LH and FSH are synthesized by the same cell, LH is packaged in dense storage granules246 with regulated secretion occurring from the basolateral surface under the control of the pulsatile secretogogue, GnRH.247–249 In contrast, FSH secretion is constitutive, linked to its synthesis, and exhibits no apparent polarity of secretion.247–250

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

The sorting determinant for the regulated secretion of LH lies in the hydrophobic heptapeptide at the carboxy terminus of LH251,252 and a single leucine residue at position 118 contributes to the sorting of LH via the regulated pathway. The heptapeptide directs the LHβ subunit to a perinuclear sub-­domain of the endoplasmic reticulum and suggests that entrance into the regulated secretory pathway is a pre-­Golgi event.253 In vivo studies have demonstrated that FSH containing heptapeptide is released via the regulated pathway similar to LH and can enhance ovulation efficiency and prolong ovarian follicle survival.254 Additionally, sulfation of LH does not play a role in its regulated secretion,255 but may increase its clearance from circulation and thereby decrease LH bioactivity.39 Although LHβ and CGβ share 85% sequence identity, hCG is not stored in granules but rather is secreted constitutively into the maternal circulation at the apical side of trophoblasts.256 The CTP sequence, unique to hCG, is the important determinant in the constitutive secretion257 and the O-­linked oligosaccharides in the CTP are essential for the apical release of hCG.258

Gonadotropin Clinical Significance • Urinary or serum hCG is used for pregnancy determination and hyperglycosylated hCG is a biomarker for pregnancy complications. • Ectopic production of hCGβ and hyperglycosylated hCG occurs in several malignancies and these molecules are used as diagnostic markers. • Recombinant gonadotropins are useful therapeutics for infertility.

Pathophysiological Expression There are reports of small amounts of hCG in the pituitary and gonadotropins in nonpituitary tissues,65,259,260 but specific physiological functions have not been ascribed to these ectopically produced hormones. In contrast, there is ample evidence supporting ectopic production in a variety of disorders. It is well known that hCG is expressed in malignant forms of gestational trophoblastic disease, e.g., invasive mole and choriocarcinoma.225,241,261,262 In men and women, hCGβ, hyperglycosylated hCGβ, and occasionally intact hCG are expressed in a variety of other malignancies, including breast, bladder, colorectal, endometrial, head and neck, hematological, lung, neuroendocrine, oral/facial, pancreatic, prostate, and testicular cancer.261,263–268 The detection of free CGβ subunit in these malignancies is generally associated with poor prognosis. In an analysis of human GCA, LHB, and CGB gene expression in breast cancer, studies showed that most normal tissues expressed only CGB7, while CGB3, CGB5, and CGB8 were expressed in trophoblastic tissues and correlated with the malignant transformation of breast cancer and other nontrophoblastic malignancies.129,269 However, human CGA, LHB, CGB1, CGB2, and CGB7 were not upregulated in breast cancer. A possible role of LH in cognition and the etiology and progression of Alzheimer disease has been postulated.270–274 Reduction in gonadal steroids and increase in serum LH that occur with aging and menopause are associated with a decline in cognition associated with Alzheimer disease and both LH and LHCGR are reported to be expressed in the brain.275–277 Of interest is the observation that LH and hCG modulate the processing of the amyloid-­β precursor protein, yielding deposition of amyloid-­β peptide.278,279 Genetic ablation of Lhcgr in amyloid precursor protein transgenic mice improved the amyloid pathology suggesting that chronic elevation of LH may promote the amyloid-­β plaque formation.280 Recent studies in rodents suggest an inverse relationship between peripheral and brain LH.274 While peripheral LH increased in ovariectomized mice, brain LH decreased. The functional and plasticity defects associated with ovariectomy could be rescued by intracerebroventricular

31

administration of hCG, suggesting that brain LH signaling plays a positive role in cognition.281

Diagnostic Applications Immunoassay-­based measurements of the serum concentrations of pituitary-­derived gonadotropins have been the mainstay of monitoring the functionality of the hypothalamic-­ pituitary-­ gonadal axis. Detection of urinary or serum hCG is a gold standard for pregnancy determination and management and for monitoring trophoblastic malignancies. Specialized biomarkers include hyperglycosylated hCG for the early detection of pregnancy or its complications235 and as a complement to detect Down syndrome pregnancies,236,282 particularly when coupled with other serum markers, α-­fetoprotein, and estradiol.283 hCGβ and hyperglycosylated hCGβ are used as tumor markers for the detection of malignancies.241,261 Immunocytochemistry is also commonly used in evaluating the expression of hCG β in suspected tumor tissue sections. It has been recognized for decades that multiple variants of hCG are present in normal pregnancy, giving rise to micro-­and macroheterogeneity.284 These include intact or heterodimeric hormone, nicked hCG, heterodimeric hormone with bond cleavages in the hCG β-­subunit 43–48 region, free α and β-­subunits, nicked hCG β-­subunit core fragment (i.e., free β-­subunit with bond cleavages in the 43–48 region), and hCG β-­subunit core fragment consisting of two disulfide-­linked fragments, 6–40 connected to 55–92.224 Hyperglycosylated hCG can present with a similar number of derivatives. Some of these variants may be useful diagnostic tools for germ cell and gynecological malignancies.241 Therefore, hCG, assays must be capable of distinguishing the variants. Several reviews, workshop proceedings, and reports have addressed this issue and the challenges of obtaining and using appropriate standards.26,240,241,262,284 The preparation and adoption of universal standards, coupled with complete characterization and disclosure of antibody specificities, will greatly facilitate the standardization of glycoprotein hormone immunoassays. While immunoreactivity is the primary technique for determining hormone concentrations in body fluids, it is often also necessary to measure bioactivity. The earlier cumbersome in vivo assays for the glycoprotein hormones have, by and large, been replaced with radioreceptor and signaling assays in transfected cells. Such measurements provide quantitative data on hormone-­ receptor binding and efficacy of signal transduction, but they give no information on circulatory half-­life and, thus, in vivo potency. For this, animal and human studies are obviously required.

Therapeutic Preparations Historically, gonadotropin therapeutic preparations were purified from urine. Those preparations were improved into highly purified preparations by further exhaustive biochemical purification procedures. The dawn of molecular biology allowed for the preparation of recombinant gonadotropins to be used in the treatment of infertility. These were expressed in Chinese hamster ovary (CHO) cells originally but recently have also been expressed in human cell lines to approximate the human glycosylation profile more closely.285 Several of these commercial preparations of recombinant gonadotropins, such as follitropin (FSH), are identical in amino acid sequence, to the naturally occurring hormones. New preparations which do not significantly differ from the originator comparator in their biochemical and pharmacodynamic effects are classified as biosimilars. This allows physicians to switch between preparations because they are interchangeable. Clinical similarity (interchangeable) must be achieved in any given patient. This is critically important since gonadotropins are administered more than once. This requires that there is no risk of alternating or

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32

PART I  The Fundamentals of Reproduction

switching between the biosimilar product and the reference product beyond the risk of using the originator product throughout. In the case of follitropin, there are several alternatives that differ from the originator preparation and/or require a different therapeutic treatment paradigm. These originated due to differences in properties despite production in CHO cells or because they have been expressed in human cell lines. In either case, such nuances of response are likely attributable to variation in glycosylation.285 Indeed, glycosylation of the naturally occurring as well as the recombinant preparations of follitropin has been exhaustively shown. This has led to the discovery of significant biological differences due to the absence or presence of glycosylation of the beta subunit of follitropin.286 Moreover, the ratios of these two forms vary during the menstrual cycle and with age.287 This has led to the notion that better follitropin therapeutics could be produced by developing treatment paradigms that implement each glycoform in a combined therapeutic approach.286 Genetically engineered amino acid substitutions have produced super-­agonists of gonadotropins.288 These could potentially decrease the cost per injection. Engineering additional glycosylation sequons increased glycosylation with an aim to increase circulatory half-­life.288 Another approach that has proven successful in extending the half-­life of FSH is an engineered extension at the α N-­terminus with two sites of N-­glycosylation.289 Improved gonadotropin therapeutics with increased half-­life have been brought to market. That involved taking advantage of the hCG β-­subunit CTP which underpins the longer circulatory half-­life of hCG compared to other gonadotropins. This approach has been utilized most effectively in producing long-­ acting analogs of FSH with a CTP engineered to the β subunit C-­terminus.290–293 This new follitropin preparation is currently available in Europe but has not been approved in the US at the time of this writing. Along these lines, one of the difficulties in the use of gonadotropins by patients at home is the difficulty and discomfort of injections. Alternative modalities of administration would be welcomed. One clever approach is to develop micronized gonadotropin preparations that can be inhaled and cross the endothelium to enter the bloodstream.294

GONADOTROPIN RECEPTORS • Gonadotropin receptors are seven-­ transmembrane domain G protein-­coupled receptors. • These receptors function as signal effectors and couplers to transducing agents resident in the cytoplasm. • Initial recognition of their cognate gonadotropin is followed by receptor activation, the transmission of intracellular signals, and regulation of receptor residence time on the cell surface/plasma membrane.    The gonadotropin receptors and the thyroid-­stimulating hormone receptor (TSHR) belong to the highly conserved subfamily of GPCRs, the so-­called Rhodopsin family, and more specifically to the δ-­group of this large class of GPCRs, according to the phylogenetic classification proposed by Fredriksson.295 GPCRs are membrane receptors that vary considerably in molecular size but that share a common molecular topology consisting of a single polypeptide chain of variable length that traverses the lipid bilayer forming seven characteristic transmembrane hydrophobic α-­helices (transmembrane domains [TMDs]) connected by alternating extracellular and intracellular sequences or loops (EL and IL, respectively), with an extracellular NH2-­terminus and an intracellular carboxyl-­ terminal domain (Ctail). These receptors characteristically bind one or several heterotrimeric G proteins that become activated upon agonist binding, which in turn act as mediators of effector (enzymes and/or ion channels) activation and intracellular signaling. In particular, the glycoprotein hormone (GPH) receptors (GPHR) are characterized by the presence of a large extracellular domain (ECD) or ectodomain

containing several leucine-­rich repeats (LRR) (Figs. 2.5 and 2.6), where recognition and high-­affinity binding of the corresponding GPHs occur.

Gonadotropin Receptor Proteins Protein Structural Attributes The human (h) LHCGR and hFSHR (http://www.ncbi.nlm.nih. gov/gene/3973, and http://www.ncbi.nlm.nih.gov/gene/2492) are 699 and 695 amino acid residues long, respectively. By convention, the primary sequences of the hLHCGR and hFSHR are numbered from the initiator methionine of their precursor sequences obtained by virtual translation of the open reading frames of the cognate cDNA,296,297 which includes the signal peptide. The most likely sites of cleavage of signal peptides are predicted to be between residues 24 and 25 of the hLHCGR and residues 17 and 18 of the hFSHR, which yields fully processed, mature proteins of 675 and 678 amino acid residues long with apparent molecular weights of 68 to 75 kDa and ∼75 kDa for the immature forms and 85 to 95 and ∼80 kDa for the mature, fully glycosylated membrane expressed forms, respectively.4,296 Thus, the predicted NH2-­terminus start sites for the hLHCGR and the hFSHR are Leu25 and Cys18 (Figs. 2.5, 2.6, and 2.10). Both the FSHR and the LHCGR exhibit a high degree of amino acid sequence homology. Whereas the ECD amino acid sequences of the gonadotropin receptors are approximately 46% identical, the 7TMD sequence portion of the receptors shares a nearly 72% homology.4,296,298–300 Among the three domains of the gonadotropin receptors, the intracellular regions exhibit the lowest amino acid sequence homology (∼27% identity), with the exception of the amino-­terminal portions of their carboxy-­ terminal tail. The ECD of the receptors is essential for hormone recognition and binding as well as for initiating receptor activation, whereas the 7TMD propagates the conformational changes induced by orthosteric agonist binding to the ECD. In turn, these conformational changes, including cytosolic regions of the receptor, promote binding and activation of downstream signaling effectors. The intracellular domains are closely related to coupling and activation of effectors as well as with receptor trafficking, agonist-­stimulated uncoupling, and desensitization.

The Extracellular Domain As mentioned above, GPHRs are characterized by a large ECD where recognition and binding of their cognate ligands occurs. This domain can be divided into three subregions: an NH2-­ terminal cysteine-­ rich region, a region composed of several copies of a structural motif rich in leucine residues (the LRR, 12 in the hFSHR, and 9 in the hLHCGR and hTSHR), which is shared with a number of other membrane receptors involved in ligand selectivity and specific protein-­protein interactions (Figs. 2.5 and 2.6),20,301,302 and a carboxyl-­terminal cysteine-­ rich domain. This latter domain displays the so-­called hinge region, which structurally links the leucine-­rich ECD with the serpentine, seven-­transmembrane domain (7TMD) of GPHRs that is involved in high-­ affinity hormone binding, receptor activation, intramolecular signal transduction, and/or silencing the basal activity of the receptor in the absence of ligand.303 In the carboxyl-­terminal end of this region, there is a particularly important amino acid sequence (FNPCEDIMGY) which behaves as an internal agonistic unit upon structural changes in the ECD provoked by hormone binding.304,305 The ECD of gonadotropin receptors contains several putative glycosylation sites, six in the hLHCGR and three in the hFSHR. The only direct biochemical evidence that exists as to which sites are glycosylated in either comes from the crystal structures of The structures show

A

2

B

C

Fig. 2.5 Schematic representation of the human FSHR structure, including the amino acid sequence (circles).  The numbering of amino acid residues includes the leader sequence. (A) Hormone binding subdomain (HBSD) of the extracellular domain. The FSHR residues that are buried in the FSH/FSHR interface and located in the high-­affinity binding site are shown in colored circles: residues shown in green, blue, or orange are buried at the receptor-­ligand interface by FSHα, FSHβ, or both subunits, respectively. β-­Strands located in the concave (corresponding to the leucine-­rich repeats) or convex surface of the HBSD are indicated by the gray arrows (Schematic according to Fan and Hendrickson structure).19 (B) Signal specificity subdomain (Hinge). The regular secondary structure elements of the leucine-­rich repeats 11 and 12 are shown as grey arrows for the β-­strands and as a cylinder for the α-­helix. The Y335 residue, which interacts with a binding pocket located in the interface of the FSH α-­ and β-­subunits (represented by the cherry and orange freeform drawings, respectively), is surrounded by a broken-­line red oval. Amino acid residues in this region of the FSHR that are buried in the FSH/FSHR interface are shown in blue (M265, which is buried by FSHβ) and grey (residues buried by FSHα and/or FSH.314 (C) Seven-­transmembrane domains (7TMDs), including the extracellular loops (EL), intracellular loops (IL), and carboxyl-­ terminal domain (Ctail). Alpha helices of the 7TMD (1 to 7) are shown as cylinders. Also indicated are amino acid residues that are involved in APPL1-­ IL1 interaction (K393) or that conform to important motifs and sequences involved in receptor function: (a) The FNPCEDIMGY sequence involved in internal agonist activity; (b) The F(X6)LL and BXXBB motifs (the latter motif found reversed at the NH2-­terminal end of the carboxy-­terminal tail) involved in receptor trafficking; (c) Motifs involved in G protein coupling and receptor activation (ERW and NPXXY motifs at the TMD3-­IL2 junction and within TMD7, respectively; and another reversed BXXBB motif in the IL3); (d) The Ser/Thr cluster for receptor phosphorylation by GRK 2 (class B cluster at the Ctail); and (e) The target cysteine residues for palmitoylation (C644, 646 and 672) 297,544,696. The locations of the naturally occurring loss-­of-­function mutations reported to date are shown as red-­colored circles, whereas those of gain-­of-­function mutations are shown as green squares. Gain-­of-­function mutations at D567, I545, and T449 lead to promiscuous activation of the receptor by other glycoprotein hormones, whereas the mutation at N431 leads to altered desensitization and internalization of the FSHR.333 Mutation at V514 (magenta circle in the EL2) led to increased plasma membrane expression of the receptor and OHSS at low FSH doses. The location of the mutation in the HBSD domain leading to promiscuous ligand binding is also shown in magenta (S128). The location of the common A307T and N680S polymorphisms are indicated in yellow, the disulfide bonds are shown as red broken lines, and the glycosylation sites are by the arbor-like structures.

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PART I  The Fundamentals of Reproduction

A

B

C

Fig. 2.6 Schematic representation of the human LHCGR structure, including the amino acid sequence (circles).  The numbering of amino acid residues includes the leader sequence. (A) Hormone binding subdomain (HBSD) of the extracellular domain. The LHCGR residues that are probably buried in the LHCG/LHCGR interface and located in the high-­affinity binding site are shown in colored circles: residues shown in green, blue, or orange are buried at the receptor-­ligand interface by the α, β, or both subunits of LH or hCG, respectively. (B) Signal specificity subdomain (Hinge). The Y331 residue, which interacts with a binding pocket located in the interface of the FSH α-­ and β-­subunits (represented by the cherry and blue freeform drawings, respectively), is surrounded by a broken-­line red oval (schematic prepared by analogy to the reported tri-­dimensional structure of the FSHR ectodomain in complex with FSH19,20,314). Also shown is the FNPCEDIMGY sequence involved in internal agonist activity. (C) Seven-­transmembrane domains (7TMDs) including the extracellular loops (EL), intracellular loops (IL), and carboxyl-­terminal domain (Ctail). Alpha helices of the 7TMD (1 to 7) are shown as cylinders. The schematics also show the location of some important motifs and sequences involved in receptor function: (a) The F(X6)LL motif involved in receptor trafficking from the endoplasmic reticulum to the plasma membrane; (b) Motifs involved in G protein coupling and receptor activation (ERW and NPXXY motifs at the TMD3-­IL2 junction and within TMD7, respectively; and the reversed BXXBB motif in the IL3); (c) The target cysteine residues for palmitoylation (C643 and C644) and residues involved in postendocytic trafficking (indicated by an asterisk). The locations of the naturally occurring loss-­of-­function mutations reported to date are shown as red-­colored circles, whereas those of gaincircles arbor-­like structures.

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

that carbohydrate is attached at residue N191, which protrudes into the solvent, whereas no carbohydrate is attached at residue N199, which protrudes from the flat β-­sheet into the hormone-­ receptor binding interface. No structural information is available for residues 293 and 318 at this time. Nevertheless, studies with the rat FSHR have suggested that the human receptor might be glycosylated at two of the three glycosylation consensus sequences at positions (191, 199, and 293), whereas in the rat LHCGR at least five, or perhaps all six of the glycosylation sites have been reported to be glycosylated.296 Although carbohydrate does not play any role in hormone binding, its role in folding and trafficking of the receptor to the cell surface will be discussed below. The ECD of gonadotropin receptors also has several disulfide bonds (5 in each receptor) that stabilize the three-­dimensional structure of the ECD, including the hinge region (Figs. 2.5 and 2.6). However, it is uncertain whether the inner Cys-­bridge in the hinge region of the hLHCGR is essential or even existent because in new world monkeys LHCGR (“LHCGR type 2”) with constitutive skipping of exon 10 (which encodes 27 amino acids in the hinge region) is still normally functioning when stimulated by CG.306 The first crystal structure of a large portion of the ECD of the hFSHR (residues 1–268, see Fig. 2.7) complexed with a single chain hFSH analog shed important light on the three-­ dimensional structure of this receptor region and its involvement in ligand binding and receptor activation.19,20 Although previous biochemical and in silico studies307,308 had predicted the relationship between the GPHR ectodomain and GPH agonist binding and recognition, it was not until the first structure of the FSH in complex with the extracellular-­hormone binding domain of the FSHR (FSHRHB) was established19 that this relationship was fully documented; this structure, however, did not include the hinge region, which had been considered as a separate structure essential for FSHR activation as had been suggested by biochemical studies.309–312 The Fan-­ Hendrickson’s structure showed that hFSH binds to hFSHRHB like a “handclasp,” as previously predicted by in silico studies.307 It also shows that most β-­strands in the inner surface participate in FSH binding, both hormone subunits are involved in the specificity of binding to the receptor with charge and stereochemical dominance dictating specificity, and that carbohydrates do not participate in the binding interface of the FSH–FSHRHB structure, but rather are sequestered to the periphery of the complex.19,313 A comparison of the crystal structure of free hFSH18 with that of the receptor-­bound hormone19 also revealed that the structures of the free and bound hFSH are quite similar but the hormone is more rigid when bound to the receptor, perhaps because the β-­sheet structure of the receptor ECD is rigid. The most obvious change in the hormone is on the carboxyl-­terminus of FSHαwhich becomes buried at the receptor interface where it forms contacts with receptor residues that are highly conserved among the three GPHRs.19 Subsequently, the crystal structure of FSH bound with the entire FSHR ectodomain (FSHRED) was determined20 (Fig. 2.8A). The structure described more clearly the role of the hFSHR ECD (and other GPHR as well) in ligand binding and, more importantly, receptor activation. The structure suggests that FSH is initially recruited by the previously described19 FSHRHB through high-­affinity interactions between the gonadotropin and the concave surface of LRRs 1 to 8. However, the interface between the FSH and the FSHRED is broader than that previously identified on the FSH-­FSHRHB structure due to the presence of secondary interaction sites (Fig. 2.8B). Accordingly binding of FSH to the FSHR hormone-­binding domain provokes conformational alterations in the L2β loop (V38β to Q48β) of FSH that lead to interactions between residues in this loop and LRRs 8 and 9, as well as to interactions of FSHR residues located in the hinge region (see below) with

35

2

Fig. 2.7 Crystal structure of deglycosylated hFSH/FSHRHB. The hormone-­binding domain of the receptor is colored magenta. The beta subunit is colored blue and the alpha subunit is colored green. Carbohydrate residues were modeled on the alpha subunit (yellow) and the beta subunit (green). The structure 1XWD was obtained from the protein database.

residues on both FSH subunits. Specificity of the FSHR for its ligand is determined by several residues including L55, E76, R101, K179, and I222, among which L55 and K179 are important to distinguish between LH, hCG, and FSH due to their interaction with the “seat belt” of FSHβ, whereas the other residues dictate specificity against TSH.20,314 A detailed map of interaction between residues from FSH and the FSHED is shown in Fig. 2.9. The FSHR ECD structure identified the hinge region [or signal specificity subdomain (SSSD)] as an integral part of the ectodomain (Fig. 2.5); the structure determined also confirmed that this particular region plays an essential role in mutation-­ provoked receptor activation, as earlier biochemical studies on the FSHR and TSHR had indicated.300,309,315–317 In fact, these and more recent biochemical studies305 strongly suggest that the ectodomain of the GPHRs acts as a tethered inverse agonist, which switches to an agonist upon ligand binding (i.e., the unoccupied hinge region is inhibitory of receptor activation) and activation of the internal FNPCEDIMGY sequence unit located in the carboxy-­terminal of the hinge region (Figs. 2.5 and 2.6). The hinge region of the hFSH bears a sulfated Tyr residue (Y335; Y331 in the hLHCGR) that interacts with a binding pocket located in the interface of the α-­ and β-­subunits of FSH and is formed via conformational changes in the ligand occurring after binding to the hormone-­ binding subdomain (Fig. 2.5). The sulfated Y335 is located right after a rigid hairpin loop which is thought to lift when Y335 binds to the α-­β pocket; this shift of the loop is thought to unlock the inhibitory effects of the loop on the 7TMD, leading to conformational changes in the latter domain and eventually to activation of the receptor. This tethered inverse agonist region in the ECD had been previously mapped to the hairpin loop segments 296 to As part of this process, a fixed short helix

36

PART I  The Fundamentals of Reproduction

V57

Glycan-N52

Y58

Y110

2 T75 T60 R62

E87

L73 E84 D71

R59

3 H68

A

B Fig. 2.8 Follicle-­stimulating hormone (FSH)-­FSHR ectodomain (FSHRED) trimer.  (A) Top view of the trimeric complex. FSH α-­and β-­subunits are shown in green and light-­grey ribbons, respectively, while FSHRED is shown in magenta. Oxygen atoms at the side chain of sulfated Y335 are represented as red-­colored balls. The carbohydrate atoms at N52α are shown as yellow balls. Inset: A zoomed region shows the detailed interactions between FSH and FSHRED at the trimer interface. (B) A theoretical model of a single fully glycosylated FSH molecule binding to an FSHR trimer, viewing from the top. For clarity, glycosylation is omitted, except for N52α of the hormone, which is located in the inner space of the FSH–FSHR trimer. The receptor trimer is shown as a magenta surface. The α chain of FSH is shown as a green ribbon, the β chain as a blue ribbon, and the carbohydrate as yellow balls. (From Jiang X, Dias JA, He X. Structural biology of glycoprotein hormones and their receptors: insights to signaling. Mol Cell Endocrinol. 2014;382:424–451, with permission from Elsevier.)

formed by residues S273 to A279 (S277 and R283 in the hLHCGR) rotates, functioning as a pivot, additionally contributing to the conformational change of the FSHR SSSD. The importance of this helix movement in FSHR activation is emphasized by the finding that substitution of the S273 and S277 residues in the hFSHR and hLHCGR, respectively, with nonpolar hydrophobic residues (e.g., S273I and S277I mutants) leads to constitutive activation of the receptors.318 In addition, the disulfide bond C275 to C346 (Fig. 2.5) (C279-­C343 in the hLHCGR) fastens the last β-­strand (LRR12) to the short helix forming a rigid body. Meanwhile, the disulfide bridge C276 to C356 (C280–C353 in the hLHCGR) ties this helix to the last few residues before the first TMD. Due to these constraints, the movement of the hairpin loop that occurs upon ligand binding could directly influence the conformation of the TMD helix 1, thereby promoting rearrangement within

the remaining TMDs, ultimately leading to receptor activation. Given the similarity among the structures of GPH and GPHR, it is highly possible that all GPHRs share this 2-­step recognition process. For example, ligand recruitment by the HBSD followed by SSSD sulfated tyrosine docking to either Y335 in the FSHR, Y331 in the hLHCGR, or Y385 in the TSHR, albeit with some differences in the spatial arrangements when hLH and hCG interact with the sulfated Y331 residue in the hLHCGR compared with hFSH interaction with Y335.319 In fact, mutation of GPHR in this critical sulfated tyrosine residue led to the loss of sensitivity to their corresponding ligands.310,311,320 Further, substitutions in amino acid residues located below the sulfated tyrosine binding pocket (αF74E) or at the potential exosite (βL73E) increased signaling of the hFSH mutants, perhaps by forcing the hairpin loop upwards at the top of the pocket.20 Although both LH and hCG bind the same receptor, differences in hLHCGR binding, activation, and signaling321,322 have been suggested on the basis of homology modeling and site-­directed mutagenesis on the hinge region, specifically in the primate-­specific exon 10,319,323 which is essential for full LHCGR activation by human LH but not by human hCG.306 In this vein, it is important to note that the carboxyl-­terminal flanking sequence of the L2β loop (which participates in hLH and hCG binding to the hinge region of the hLHCGR) differs between hLH and hCG with respect to proline residues, leading to differences in the interaction between hLH and hCG with the sulfated Y331.319 The crystal structure of the large region of the ECD of the TSHR in a complex with a TSHR autoantibody has also been solved.324 Although the number of LRRs is different between the TSHR and the FSHR, the overall structure of their corresponding ECDs is very similar.325 Interestingly, the TSHR surface that binds the autoantibody is remarkably similar to the surface of the FSHR that binds FSH.324,326 An interesting nuance is hFSHR (and hTSHR as well, but not hLHCGR) promiscuity for ligand specificity occurring upon structural modifications in the ECD caused by particular mutations (Figs. 2.5). This phenomenon may convey important consequences in the clinical arena. GPH-­GPHR pairs have evolved in such a manner that a limited number of residues in both the “seat-­belt” domain of the ligand and the LRRs of the receptor at the hormone-­binding domain participate in electrostatic interactions at the receptor-­hormone interface to define binding and specificity. Due to the structural similarities between the GPH and the GPHR, it is conceivable that “cross-­ activation” of a given GPHR by other than its cognate ligand may occur, albeit with a low binding affinity, without triggering basal detectable receptor activation in physiological conditions. Therefore, it is not surprising that substitutions in key residues that directly or indirectly participate in the interaction of the receptor with its cognate ligand may decrease structurally-­related ligand discrimination resulting in the interaction of the altered receptor with other than its own cognate GPH. For example, this is the case of the S128Y mutation at the hFSHR (Fig. 2.5), which leads to pregnancy-­ associated ovarian hyperstimulation syndrome (OHSS).327 In its severe form, OHSS may be life-­threatening due to increased responsiveness of the FSHR to hCG, which circulates at very high levels during the first trimester of pregnancy.328 In this particular mutation, the S→Y replacement allows the hFSHR to hydrogen bond αR95 of hCG, leading to receptor activation. A different panorama is observed in the case of mutations in the 7TMD leading to constitutive activation of the FSHR and concomitantly promiscuous binding of hCG and/or TSH, as will be discussed below.

The Extracellular Loops (EL) The extracellular loops or exoloops of GPHRs transduce the signal generated by the ligand-ectodomain interactions to the transmembrane helices either through direct hormonal contact

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

37

2 Y103

LRR1 L55

L99

LRR2 K45

D81

LRR3 LRR4

K104

V96 D93

Y124

D90

LRR5 D153

LRR6 LRR7

R42

K51 R97

E76 R101

Y88 S89

K91 D150

K179

LRR8 I222

LRR9 P45

LRR10

FSHRHB

FSH

Fig. 2.9 Schematic illustration of detailed interaction at the FSH–FSHR interface.  Contacting residues from FSHRHB are shown as yellow dots and those from FSHα as red dots and FSHβ as blue dots. The middle column summarizes the specific side-­chain interactions between FSH and FSHRHB. Interactions that contribute to common affinities among all the GPH–GPHR family members are shown as green-­filled circles (for charge– charge interactions) or boxes (for non-­charged atomic contacts) and they are connected by green lines back to the yellow dots in FSHR or red or blue dots in FSH α-­ or β-­subunits, respectively. Interactions that contribute to specificity are shown as purple-­filled circles or boxes and they are connected by purple lines to the dotted residues in FSHR and FSH. (Reproduced from Jiang X, Dias JA, He X. Structural biology of glycoprotein hormones and their receptors: insights to signaling. Mol Cell Endocrinol. 2014;382(1):424–451. with permission from Elsevier.)

and/or by modulating the interdomain interactions between the hinge region and the TMD. In fact, EL1 and 3 are solvent exposed and accessible to gonadotropin hormone, representing potential secondary binding sites for the gonadotropin, specifically at its α-­subunit tips in the bound state as supported by structural studies on the FSHR.329,330 Moreover, according to mutagenic and functional studies and the 7TMD theoretical model of the hFSHR proposed by Jiang et al,314 it seems that the ELs interact with the hairpin loop of the hinge region to trigger FSHR activation; apparently, lifting of the sulfated Y335 to the FSHα/β binding pocket frees the hinge-­tethered exoloops, releasing the inhibitory influence of the ectodomain on receptor activation.314 Accordingly, mutations in residues located at the hFSHR and hLHCGR ELs 1 to 3, besides altering the intracellular traffic of the receptor,331–333 may also attenuate agonist binding,334–336 alter hormone-­ stimulated signal transduction,331,332,335–340 or provoke constitutive activation of the receptor.333 The relationship between the exoloops of the gonadotropin receptors and the hinge region is further supported by studies in the TSHR, in which particular residues (e.g., Y563 and K565 at the exoloop 2 (Y511 and K513 in the FSHR; and Y508 and K510 in the LHCGR]) are crucial for ligand-­stimulated receptor activation.299,341

The Transmembrane Domains As described above, the α helices forming the 7TMD of the gonadotropin receptors are joined by three alternating

intracellular (IL) and extracellular (EL) loops (see Figs. 2.5 and 2.6). Although a three-­dimensional structure of the 7TMD of the GPHRs is lacking, the three-­dimensional structures of a number of other GPCRs with short ECDs have now been solved342 (also see gpcr.usc.edu) and the TMD of the gonadotropin receptors is likely to be very similar, particularly among the rhodopsin/β2-­ adrenergic receptor-­like subfamily of GPCRs. TMD residues that are highly conserved among this subfamily of GPCRs are highlighted in Fig. 2.10. Both gonadotropin receptors exhibit the general ERW and NPXXY motifs (at the TMD3-­IL2 junction and within TMD7, respectively), which are common features of the rhodopsin-­like GPCR subfamily and play a crucial role in receptor activation. Further, the importance of the conformational changes on the 7TMD in receptor activation is emphasized by the fact that the majority of natural mutations leading to constitutive activation of the GPHRs are located in this receptor domain (Figs. 2.5 and 2.6).343 In addition, several amino acid residues in the cytoplasmic face of the 7TMD of gonadotropin receptors (i.e., at the ILs) also play important roles as interactors with effectors and adapters involved in intracellular signaling and postendocytic processing (see below).344–349 Through homology modeling based on biophysical, structural, and computational modeling data from other GPCRs it has been possible to explore the molecular mechanisms subserving receptor activation and propagation of the activation signal from the 7TMD to the intracellular domains. In particular, three structures proved particularly crucial to understanding the free form of opsin cocrystallized

38

PART I  The Fundamentals of Reproduction N-terminal cysteine-rich region

hLHR 25 L R E A L C C hFSHR 18

- E P C N C V P D G - - - - - - - A L R C P G - P T A G L T R L S L A Y L P V K VI P S Q A F R G L N E V I K I E I S Q I D S 85 H R I C H C S N R V F LC Q E S K V T E 1 P S D L P R N A I E L R F V L T K L R VI Q K G A F S G F G D L E K I E I S Q N D V 82

hLHR 86 L E R I E A N A F D N L L N L S E I L I Q N T K N L R Y I E P G A F I N L P G L K Y L S I C N T G I R K F P D V T K V F S S E S N F I L E I 155 hFSHR 83 L E V I E A D V F S N L P K L H E I R I E K A N N L L Y I N P E A F Q N L P N L Q Y L L I S N T G I K H L P D V H K I H S - L Q K V L L D I 151 Leucine-rich motif region hLHR 156 C D N L H I T T I P G N A F O G M N N E S V T L K L Y G N G F E E V Q S H A F N G T T L T S L E L K E N V H L E K M H N G A F R G A T G P K 255 hFSHR 152 Q D N I N I H T I E R N S F V G L S F E S V I L WL N K N G I Q E I H N C A F N G TQ L D E L N L S D N N N L E E L P N D V F H G A S G P V 221

hLHR 226 T L D I S S T K L Q A L P S Y G L E S I Q R L I A T S S Y S L K K L P S R E T F V N L L E A T L T Y P S H C C A F R N L P T K E O N F S H S 295 hFSHR 222 I L D I S R T R I H S L P S Y G L E N L K K L R A R S T Y N L K K L P N L E K L V A L M E A S L T Y P S H C C A F A N W R R Q I S E L H P I 291 Hinge region hLHR 296 I S E N F S K Q C E S T V R K V N N K T L Y S S M L A E S - - - - - - E L S G W DY E Y G F C L P K T P R - C A P E P D A F N P C E D I M G 358 hFSHR 292 C N K S I L R Q E V D Y M T Q T R G Q R S S L A E D N E S S Y S R G F D MT Y T E F D Y D L C N E V V D V T C S P K P D A F N P C E D I M G 361 IL-1

TM-1

TM-2

EL-1

hLHR 359 Y D F L R V L I W L I N I L A I M G N M T V L F V L L T S R Y K L T V P R F L M C N L S F A D F C M G L Y L L L I A S V D S Q T K G Q Y hFSHR 362 Y N I L R V L I W F I S I L A I T G N I I V L V I L T T S QY K L T V P R F L M C N L A F A D L C I G I Y L L L I A S V D I H T K S Q Y

IL-2

TM-3

N 428 N 431

TM-4

hLHR 429 H A I D W Q T G S G C S T A G F F T V F A S E L S V Y T L T V I T L E R W H T I T Y A I H L D Q K L R L R H A I L I M L G G W L F S S L I A 498 hFSHR 432 Y A I D W Q T G A G C D A A G F F T V F A S E L S V Y T L T A I T L E R W H T I T H A M Q L D C K V Q L R H A A S V M V M G W I F A F A A A 501 EL-2

IL-3

TM-5

hLHR 499 M L P L V G V S N Y M K V S I C F P M D V E T T L S Q V Y I L T I L I L N V V A F F I I C A C Y I K I Y F A V R N P E L M A T N K D T K I A 568 hFSHR 502 L F P I F G I S S Y M K V S I C L P M D I D S P L S Q L Y V M S L L V L N V L A F VV I C G C Y I H I Y L T V R N P N I V S S S S D T R I A 571 TM-6

EL-3

TM-7

Helix 8

hLHR 569 K K M A I L I F T D F T C M A P I S F F A I S A A F K V P L I T V T N S K V L L V L F Y P I N S C A N P F LY A I F T K T F Q R D F F LL L 638 hFSHR 572 K R M A M L I F T D F L C M A P I S F F A I S A S L K V P L I T V S K A K I L L V L F H P I N S C A N P F LY A I F T K N F R R D F F I L L 641

hLHR 639 S K F G C C K R R A E L Y R R K D F S A Y T S N C K N G F T G S N K P S Q S T L K L S T L H C Q G T A L L D K T hFSHR 642 S K C G C Y E M Q A Q I Y R - - T E T S S T V H N T H P R N G H C S S A P R V T S GS T Y I L V P L S H L A Q N R Y T E C

Fig. 2.10 Amino acid sequence alignment of the human luteinizing hormone/choriogonadotropin receptor (hLHCGR) and human follicle-­stimulating hormone receptor (hFSHR).  Amino acid sequences were obtained from a public website (www.ensembl.org/index.html). The boundaries of the three distinct regions of the extracellular domain discussed in the text (N-­terminal cysteine-­rich region, leucine-­rich motif region, and hinge region) are marked with green, red, and green arrows, respectively. The seven transmembrane (TM) helices and the putative cytoplasmic helix 8 are delineated by black rectangle boxes and labeled TM-­1 through TM-­7 and helix 8, respectively. The three extracellular (EL) and four intracellular loops (IL) that connect the transmembrane regions are labeled EL-­1 through EL-­3 and IL-­1 through IL-­4, respectively. Identical residues between the two receptors are shown with blue boxes. The consensus sequences for N-­linked glycosylation are shown with turquoise boxes. The tyrosine that participates in dimer formation for the FSHR and is conserved in the hLHCGR is shown in yellow. The conserved tyrosines that may be sulfated are shown in pink. The conserved cysteines that are believed to be palmitoylated are shown in the green box. Residues that are highly conserved among the rhodopsin/β2-­adrenergic family of G-­protein–coupled receptors are shown in red. (Copyright © 1999-­2008 The European Bioinformatics Institute and Genome Research Limited, and others. All rights reserved.)

with the carboxyl-­terminus of the α-­subunit of the heterotrimeric visual Gt protein350; an agonist-­ bound β2-­adrenergic receptor (AR) stabilized in the active conformation by a nanobody mimicking a G protein;351 and an agonist-­bound β2-­AR cocrystallized with heterotrimeric Gs protein (Gα β γ ).352 Based on these structures, agonist-

to lead to a set of common structural rearrangements.353 First, the extracellular part of the transmembrane bundle is initially affected by the agonist-­induced local structural changes: a. a small distortion of TMD5; b. relocation of TMD3 and TMD7; and c. reorganization of TMD5 and TMD6. Concurrently, a rearrangement of a cluster of conserved hydrophobic and

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

39

Gonadotropin Receptor Gene Expression and Regulation

aromatic residues called a “transmission switch” deeper in the receptor core occurs (which involves residues 6.48, 6.44, 5.50, 5.51, and 3.40*; generic amino acid numbering according to the Ballesteros and Weinstein nomenclature354), leading to rearrangement at the TMD3–TMD5 interface, and formation of new noncovalent contacts at the TMD5–TMD6 interface.355 Many of the residues involved in this transmission switch are highly conserved in the Rhodopsin family of GPCRs, suggesting that they are likely to constitute a common feature of GPCR activation and are now discussed in the context of the FSHR primary structure. Thus local changes would be translated into larger-­scale helical conformational changes occurring at the cytosolic side,353 resulting in rearrangements of TMD5 at its cytoplasmic side356 associated with a modification of the TMD5–TMD6 interface yielding the large-­scale relocation of the cytoplasmic side of TMD6.357 As a result, the cleft required for binding of the Gα subunit opens. Residues from the IL2 and the cytoplasmic end of TMD3 (i.e., the arginine residue of the conserved ERW sequence in the gonadotropin receptors (Figs. 2.5 and 2.6) participate in the interaction with the G protein after activation.351 Specifically, as a consequence of receptor activation, the salt bridge between residue 3.50 (which corresponds to Arg467 and Arg464 in the hFSHR and hLHCGR, respectively) and residue 6.30 (Asp567 and Asp564 in the TMD6 of hFSHR and hLHCGR, respectively) present in the inactive state would be broken.350 These biophysical and structural studies indicate that agonist binding alone may not be sufficient to stabilize fully active states of the receptor, and thus, binding of an effector protein on the cytosolic side of the receptor is necessary to reach the fully active state.358 In addition, there may not be a single active state and different ligands and allosteric modulators may stabilize distinctly different conformations, thereby giving rise to diverse downstream responses that may differ in magnitude.297,359,360

The human lutropin and follitropin receptors are encoded by single genes located in the short arm of chromosome 2 (LHCGR, http://www.ncbi.nlm.nih.gov/gene/3973, and FSHR, http://www.ncbi.nlm.nih.gov/gene/2492). The FSHR is about 54 Kb long and is composed of 10 exons, whereas the LHCGR is about 70 Kb in length and is composed of 11 exons. The first 9 exons of the FSHR and the first 10 exons of the LHCGR encode for the large ECD of the receptors, including part of the hinge region, while the carboxyl-­terminal end of the latter, the 7TMD, and the Ctail of the receptors are encoded by the large exons 10 and 11 of the FSHR and the LHCGR, respectively. The structural, molecular, and genomic similarities among both the gonadotropic hormones and their receptors strongly suggest that the gonadotropin receptor genes evolved from duplication of a common ancestral gene. In fact, the location of the genes is separated by only ∼200 Kb. Further, the similarities among the genes of the gonadotropin receptors and other GPCRs also suggest that the precursor for the gonadotropin receptor genes arose from combining a common GPCR ancestor.371,372,394,418

The Carboxyl-­Terminus (Ctail)

Transcriptional Regulation

The Ctail of the gonadotropin receptors is the most divergent of the three receptor domains. These domains host important sequences and motifs which play crucial roles in receptor function. For example, cysteine residues (Cys644 and 646 in the hFSHR and Cys 643 and 644 in the hLHCGR) are closely associated with the plasma membrane (PM) and palmitoylated.361–363 A primary sequence motif [Phe(X)6Leu-­Leu] at the NH2-­end of the Ctail lies within a helical segment which in other GPCRs is referred to as helix 8 (see gpcr.usc.edu). This sequence motif regulates the upward trafficking of the receptor from the endoplasmic reticulum to the PM.296,348,364 Additionally, a cluster of five serine and threonine residues at positions (656 and 658–661) are targets of G protein receptor kinases (GRK)365–367 that phosphorylate the hFSHR at those sites. These posttranslational modifications promote β-­arrestin recruitment. This cytoplasmic protein is a scaffold that regulates ligand-­stimulated G protein uncoupling and receptor desensitization, internalization, and intracellular signaling mediated by ERK1/2.365 Finally, Ctail residues Pro688 and Leu689 in the hFSHR and Gly687 and Thr688 in hLHCGR are involved in postendocytic trafficking of the internalized receptors.368–370 The importance of these sequences in gonadotropin receptor function briefly described above is more extensively discussed below.

The 5′ flanking regions of the LHCGR and FSHR are rich in GC nucleotides and the proximal promoters of both genes are devoid of TATA boxes and have multiple transcription initiation sites within a region similar to an initiator element. Despite these similarities, the particular and divergent functional characteristics of key regulatory features present in the LHCGR/Lhcgr promoter and the FSHR/Fshr distal regulatory sites emphasize the differential regulation of each receptor gene.372,373 Studies on the transcriptional control of the FSHR gene have been done mostly with rodent genes.373 Remarkable features of the control of FSHR gene expression include its regulation by a combination of transcriptional and posttranscriptional mechanisms. Both FSH and activin in the ovary and FSH in the testis play a role in this regard. Expression of the FSHR/Fshr requires regulatory elements located both within and outside the promoter region (i.e. 97kb upstream of exon 1 to 57kb downstream of exon 10). Within the promoter region are elements that bind NR5A1 (SF1) as well as upstream stimulatory factors (USFs). External to the promoter region, DHS#3 (a DNase I hypersensitive site) is located ∼4 kb downstream in intron 1 and is the target of the GATA-­binding protein complex and POU2F1 (octamer-­binding protein 1). Because the myoid cells employed in these studies do not express FSHR, it has been suggested that the DHS#3 element is associated with gene silencing.374 In several animal species, including humans, the FSHR promoter shows a common E box element, which contributes significantly to promoter activity, and binds its cognate USF1 and USF2 transcription factors, which regulate differential gene expression in Sertoli and granulosa cells.375–377 Studies of the Fshr have also shown that USF binding to the E box increases during differentiation and decreases, concomitantly with promoter

* The first number corresponds to the TMD where the residue is located and the second to the most conserved residue in this TMD, which is arbitrarily assigned to 50, with numbers decreasing toward NH -terminus and increasing toward carboxyl-terminus.

• Production of a biologically active receptor begins with the induction of gene expression and generation of mRNA. • Correct folding and posttranslational processing of the receptor protein are essential for trafficking of the newly synthesized receptor to the plasma membrane as well as for receptor-­receptor association. • Receptor activation is followed by signal transduction, desensitization of the receptor, internalization, a second wave of signaling, and degradation or recycling of the receptor molecule.

Genes

2

40

PART I  The Fundamentals of Reproduction

activity, upon FSH administration. This occurs because of increased expression of the DNA inhibitor of DNA binding/ differentiation-­2 protein (ID2), which inhibits E box binding and promoter activity.378–380 In some species, including rodents, humans, and sheep, the FSHR gene promoter contains a putative estrogen response element (ERE) as well as a cAMP response element (CRE)-­like sequence, which may be involved in cAMP-­mediated transcriptional regulation.381–384 Some rodent species also contain putative cis-­acting elements for transcription factors of the HMG-­box SRY/Sox family, which seem important in the positive transcriptional regulation of the FSHR gene.383 Nevertheless, many of the regulatory factors extending beyond the FSHR gene still remain unknown. This was evident when transgenic mice carrying a yeast artificial chromosome (YAC) with the entire Fshr plus 50 kbs 5′ and 30 kbs 3′ bordering sequences failed to express the receptor in Sertoli cells.375 The cis-­acting elements and trans-­acting factors that control the transcription of the mouse, rat, and human LHCGR have been examined in more detail.372,385 The major transcriptional sites of the TATA-­less LHCGR gene promoter are located within a 173 to 176 bp domain; although in the rat, transcription of the LHCGR gene is inhibited by upstream sequences located from -­175 bp and -­2056 bp, in the human promoter this inhibition is minimal.386 Elements regulating transcriptional activation of the LHCGR gene include an imperfect ERE/DR motif, as well as SP1(II) and SPi(I) elements located at -­79 bp and -­119 bp from the codon initiation site (ATG) in the human gene.387 These elements and their corresponding transcription factors either inhibit or stimulate transcription depending on the regulators bound (EAR2/EAR3 or TR4).388 The ERE/DR motif binds the nuclear orphan receptors (OR) factors EAR2 and EAR3, which inhibit transcription, and TR4, which stimulates it. EAR2/EAR3 inhibits LHCGR transcription by disrupting the positive interaction between TFIIB-­bound to the preinitiation complex and Sp1/ Sp3 factors bound to their corresponding elements. Meanwhile, Sp1 and Sp3 bound to the Sp1(I) element directly or indirectly interacts with silencing regulatory complex mSim3A/RbAp48 to repress transcription. Active silencing of LHCGR expression proceeds through crosstalk among COUP-­TF1/EARs, Sp1/Sp3, and TFIIB.388 During gonadotropin stimulation, the concentrations of ER2/ER3 change, which correlates with derepression of promoter activity. In the ovarian follicle, the derepression-­silencing cycle governed by the orphan factors depends on the stage of follicular maturation, with repression present in small, early follicles and derepression-­activation in mid-­to preovulatory follicles and luteal follicles. FSH stimulation in granulosa cells induces CYP19A1 expression and LHCGR gene transcription and expression levels necessary for the final steps in the maturation process of the oocyte and for follicular rupture. These orphans, including TR4, also bind the human DR element LHCGR in the testis LHCGR in the human. Epigenetic regulation also has been involved in LHCGR expression, with acetylation and demethylation of histones playing important roles.389,390 Acetylation occurs through inhibition of HDAC1/2 (histone deacetylase) bound to the mSin3A/RbAP48 complex or the Sp1 factor at the Sp1 (I) site, and recruitment of acetylated H3 and H4 to the promoter.391 DNA demethylation of the promoter seems necessary to overcome the inhibitory constraint imposed by the HDAC/ mSin3A complex. Thus, epigenetic silencing and activation of the LHCGR gene are achieved through the concerted action at both histone and DNA levels, with coactivators and corepressor factors playing important regulatory roles.385,392 Several signaling pathways are involved in the complex regulation of the LHCGR gene, including the PI3/PKCζ pathway, which phosphorylates Sp1 and derecruits corepressor factors, and the PKC-­ α/ERK pathway, which also phosphorylates Sp1 leading to dissociation

of inhibitor complexes such as HDAC1/mSin3A and recruitment of TFIIB.385,390,393

Transcripts Multiple transcripts of the gonadotropin receptors have been found, which apparently arise from alternative splicing, different transcriptional start sites, or the use of different polyadenylation sites.394 The ontogeny of the multiple FSHR and LHCGR transcripts in female and male rats has been reviewed elsewhere.4 Although there is some controversial evidence regarding the number and nature of multiple FSHR transcripts,373 at least 4 and 2 FSHR mRNA transcripts ranging in size from 7.7 to 1.8 kb have been detected in rat ovaries and testis, respectively, with the 2.5 kb transcript being the most abundant.395,396 Several heterogeneous FSHR splice variants have been detected in various animal species,384,397–403 including humans.404–410 Four splicing products affecting the ectodomain of the hFSHR have been detected in cumulus cells surrounding oocytes isolated from follicular aspirates of women undergoing in vitro fertilization. When these variants were expressed in vitro, a reduced response to exogenous FSH stimulation was observed.404 There also are reports of transcripts in the sheep FSHR that arise from alternative splicing and that are presumably translated into functional proteins with distinct signaling properties.411 Since many of the alternative transcripts detected are not translated into a protein or translation yields severely misfolded proteins unable to traffic to the cell surface PM, the function and regulatory role of FSHR transcript variants have been scarcely studied. Two alternately spliced transcripts of the ovine FSHR gene with altered exon 10 are particularly interesting because of their potential functional consequences.384,412–415 The first variant is similar to the full-­length wild-­type (WT) receptor except that differential splicing leads to divergence in protein sequence in the carboxyl-­terminus and that the variant is 25 amino acids shorter; this FSHR variant is expressed at the PM, fails to signal via the cAMP/PKA pathway and behaves as a dominant-­negative receptor when coexpressed with the WT receptor in HEK293 cells.414 The second FSHR variant consists of a transcript encoding only exons 1–8 along with a single putative TMD unlike anything in FSHR and a carboxyl-­terminal extension, all more characteristic of cytokine/growth factor receptors.384 When expressed in granulosa cells, the growth-­promoting effects of this variant were independent of the cAMP/PKA pathway and were rather mediated by activation of Ca2+ and ERK1/2-­dependent pathways.413 Further studies demonstrated that this particular variant is also expressed in the mouse ovary and that it is upregulated by gonadotropin stimulation.397 More recent studies in ewes416 found that the FSH-­ growth factor-­like receptor was more highly expressed than the WT variant in all-­size follicles, particularly in medium-­size follicles obtained during the beginning of estrus, suggesting that this receptor isoform may participate in follicle development in this particular animal species. Additional studies are still necessary to more precisely define the role of these and other FSHR transcript variants in mediating the pleiotropic effects of FSH (see below), particularly in humans. The transcript sizes of the LHCGR gene vary depending on the species studied. The rat receptor expresses 4 transcripts ranging in size from 6.7 to 1.8 kb, whereas in human granulosa cells two mRNA transcripts have been detected.372,417,418 Several LHCGR mRNA splice variants have been identified in different animal species,419–421 including humans.422–426 A variant lacking exon 9 isolated from a human corpus luteum is particularly interesting since it exerts dominant-­negative effects on both hFSHR and hLHCGR,424,425,427 which may represent a mechanism that regulates the gonadotropic stimulus in this ovarian structure. A cryptic exon (exon 6A) localized between exons 6 and 7 identified in primates, produces transcripts leading to nonsense-­mediated

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

mRNA decay; although the physiological significance of these variants is unknown, mutations in this cryptic exon have been associated with male pseudohermaphroditism secondary to Leydig cell hypoplasia.428,429 Recent studies suggest that gonadotropin receptors may be expressed in a number of extragonadal tissues where they could exert distinct roles in certain physiological and/or pathological conditions. This interesting aspect is further discussed below.

Posttranscriptional Regulation Posttranscriptional regulation is also an important aspect of the regulation of the LHCGR mRNA and the LHCGR levels during the preovulatory LH surge or in response to supra-­physiological levels of the gonadotropin.430 This important level of regulation seems to be mediated by several factors including an LHCGR mRNA binding protein (LRBP) which has been identified as mevalonate kinase, an enzyme that is involved in cholesterol metabolism that is also an LHCGR mRNA binding protein.430 LHCG-­ induced activation of steroidogenesis and subsequent depletion of cholesterol triggers an increase in the transcription of genes that participate in cholesterol biosynthesis, including mevalonate kinase, through a PKA/ERK1/2 signaling pathway.430,431 The increased levels of this protein may thus serve the dual role of enhancing cholesterol synthesis (an enzymatic function) and decreasing the levels of LHCGR mRNA by virtue of its ability to bind the LHCGR mRNA and promote its degradation, transiently downregulating receptor expression.432,433 This transient decrease in LHCGR mRNA levels is then followed by a full recovery and increased expression during corpus luteum development, only to eventually fall again with the regression of this highly steroidogenic ovarian structure. More recent studies have shown that levels of LRBP are regulated by the microRNA miR-­122, which activates the sterol regulatory element-­binding protein (SREBP) pathway leading to LHCG-­stimulated, PKA/ ERK1/2-­mediated upregulation of mRNA and protein LRBP levels.432,434,435 Another miRNA involved in the posttranscriptional regulation of mRNA LHCGR is miR-­ 513a-­ 3p, which interacts with the mRNA through three binding sites located in the 3′UTR region, leading to downregulation of mRNA LHCGR expression levels in human granulosa cells,436 as well as MiR-­136-­3p which has been found associated with downregulation of the LHCGR in rat granulosa cells.437 Posttranscriptional regulation of the FSHR expression has been less extensively studied, although it is well known that several factors, including TGFβ and activin increase the stability and half-­ life of the FSHR mRNA, upregulating FSHR expression.438,439 In female rats, FSH stimulation may up and downregulate mRNA and protein FSHR expression, depending on the stage of follicular maturation.395 Apart from miR-­126, which was recently shown to downregulate FSHR mRNA and to act synergistically with androgens to inhibit FSHR protein in porcine granulosa cells,440 little is known about whether other miRNAs are involved in the posttranscriptional regulation of the FSHR mRNA.

Folding, Maturation, and Intracellular Trafficking The life cycle of GPCRs begins at the endoplasmic reticulum (ER), where synthesis, folding, and assembly of proteins occur (Fig. 2.11). Properly folded receptors are then targeted to the ER-­ Golgi intermediate complex and thereafter to the Golgi apparatus and trans-­Golgi network; here, the processing is completed and the receptor proteins are ready to complete their outward trafficking to the cell surface PM, where they become accessible to their cognate ligands.441 Interaction between GPCRs and agonists at the PM triggers downward trafficking of the receptor through a series of distinct processes including (a) phos phorylation (which terminates G protein-

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and β-­arrestin recruitment, which interact with clathrin and the clathrin adaptor AP2 to drive receptor internalization into endosomes, and (b) either recycling of the receptor back to the PM or targeting to the lysosomes and/or proteasomes for degradation. Thus, the balance between trafficking from the ER to the PM and the endocytosis-­recycling/degradation pathway determines the net amount of receptor protein available to interact with agonists and elicit a measurable biological response. GPCRs are subjected to a strict quality control system (QCS) that monitors, and corrects when necessary, the folding of the nascent receptor into a three-­dimensional structure.442 By monitoring the structural and conformational features of newly synthesized proteins, the QCS determines which proteins must be retained at the ER and eventually degraded or routed to the Golgi apparatus and thereafter to the PM.443 Thus, the QCS prevents the accumulation of misfolded proteins that may aggregate and interfere with cell function. GPCR export from the ER to the Golgi is modulated by the interaction of the trafficking proteins with specialized folding factors, escort proteins, retention factors, enzymes, and members of the molecular chaperone families, which belong to the ER QCS and the so-­called proteostasis network.444–446 In particular, molecular chaperones are essential components of the ER QCS that evaluate native receptor conformation and promote delivery from the ER to the Golgi where the protein molecule is processed before final delivery of the mature form to the PM.447–450 Molecular chaperones are an important quality control mechanism that not only recognizes but also retains and targets misfolded, nonnative protein conformers for their eventual degradation via the polyubiquitination/proteasome pathway.451–454 Similar to other GPCRs, the gonadotropin receptors have to be correctly folded into a conformation to pass the QCS and be compatible with ER export442 ( Fig. 2.11). Glycosylation, which together with disulfide bonds is a frequent feature of GPCRs, occurs during biosynthesis and facilitates the folding of protein precursors by increasing their solubility and stabilizing protein conformation.455 In fact, mutations that affect glycosylation at the ECD or the formation of disulfide bonds hamper the folding process of these receptors leading to intracellular retention of the misfolded intermediates, and eventually to disease. Thus, glycosylation plays an important role not only in folding but also in the maturation and intracellular trafficking of the receptors from the ER to the cell surface PM. Studies in the rat have shown that at least one glycosylation site at the FSHR ECD is required for receptor folding and efficient trafficking to the PM.456 Lack of glycosylation of the mature rat FSHR does not affect binding or affinity, thus indicating that this structure does not participate in ligand interaction. Likewise, mutants of rat LHCGR that prevented glycosylation at the first three consensus sequences did not affect receptor synthesis or ligand binding but decreased the efficiency of receptor folding, leading to reduced maturation and increased degradation of the precursor protein.457 Thus, carbohydrates in both gonadotropin receptors, although not involved in hormone binding, are important players in the maturation process of the newly synthesized receptors, promoting their folding, conformational stability, and trafficking to the PM. Coimmunoprecipitation studies have identified some interacting proteins that support the folding of the gonadotropin receptors during their residency at the ER. These studies have shown that the folding process of the rat FSHR and LHCGR precursors (cotranslationally glycosylated at the ER by the addition of the core sugars N-­acetyl glucosamine, mannose, and glucose) involves interactions with the chaperones calnexin and calreticulin, which facilitate proper folding of intermediate glycoprotein molecules.458,459 The cycle of these chaperones predominantly centers on substrate N-­glycans present on the glycoprotein receptor precursor, adding hydrophobicity to the folding protein. The calnexin/calreticulin cycle depends on the concerted action of carbohydrate-­modifying enzymes (glycosidases I and II), which lead to the formation of

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PART I  The Fundamentals of Reproduction Ligand

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Fig. 2.11 Intracellular traffic of gonadotropin receptors from the endoplasmic reticulum (ER) to the cell surface plasma membrane.  Newly synthesized proteins fold in the ER. Correctly folded proteins are then translocated to the Golgi apparatus to initiate/complete processing such as glycosylation (magnifiers) (steps 1 and 3). Misfolded and misassembled products are retained in the ER and exposed to resident chaperones who attempt to correct folding and stabilize the protein in a conformation compatible with ER export. When correct folding fails, the misfolded protein is dislocated into the cytoplasm for proteasomal degradation (step 2). Mature receptors are then exported to the plasma membrane (step 4), where they interact with cognate ligands (steps 5 and 6). Agonist-­provoked activation of the receptor (step 6) is followed by phosphorylation of the receptor and recruitment of β-­arrestins (purple circles), as it is in the case of the FSHR, which promotes endocytosis (step 7) and internalization of the receptor (step 8). The receptor embedded in clathrin-­coated vesicles may be either targeted to lysosomes and/or proteasomes for degradation (step 9) or recycled to the plasma membrane (step 10) to interact again with cognate agonist (step 5) (see also Fig. 2.12). (Modified from Ulloa-­Aguirre A, Zarinan T, Dias JA, Conn PM. Mutations in G protein-­coupled receptors that impact receptor trafficking and reproductive function. Mol Cell Endocrinol. 2014;382(1):411–423. with permission from Elsevier.)

monoglycosylated oligosaccharide structures that interact with the chaperones as well as the removal of the remaining glucose residue from the oligosaccharide, terminating the association with the chaperones. Then the GPHR (already in its native conformation) is exported to the Golgi complex to continue processing the oligosaccharide chains. Another GPHR interacting chaperone is the protein disulfide isomerase PDI,458 which is an ER-­resident enzyme involved in disulfide bond formation of folding intermediates, and that probably acts as a cochaperone with calnexin and cal reticulin during their association with the GPHR. No interaction

of the GPHR with other molecular chaperones has been yet documented, with the exception of the TSHR. It apparently interacts with BiP, a chaperone that maintains proteins in a state competent for subsequent folding and oligomerization, and that mediates retrograde translocation of misfolded conformers for proteasomal degradation.442,460,461 Several sequence motifs present in the GPCRs are involved in the exit of the receptors from the ER and the Golgi. Among these motifs is the above mentioned F(X) LL sequence identified in the Ctail of several GPCRs, including the gonadotropin

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

receptors (Figs. 2.5 and 2.6).364 In the hFSHR, this export motif is located between amino acid residues 633 and, 641 whereas in the hLHCGR, this motif is located between residues 630 and 638.4,296 The Ctail peptide of the hFSHR also contains the minimal BBXXB motif reversed in its juxtamembrane region (residues 631–635)348; the last two residues of this motif (R634 and R635) and the preceding F633 constitute the amine-­terminal end of the highly conserved F(X)6LL motif, and thus mutations in these residues impair receptor trafficking and PM localization of the receptor.348,462 The IL3 of the hFSH and hLHCG receptors also contains this BXXBB motif (residues 569–573 in the hFSHR and 566–570 in the hLHCGR) and either deletion or replacement of the basic residues of this motif with alanine impairs PM expression of the modified receptors.348,349 Another motif that influences gonadotropin receptor folding and trafficking to the cell surface PM is the AFNGT motif (amino acid residues 193–197 in the hLHCGR and 189–193 in the hFSHR), which bears a potential glycosylation site (N195GT and N191GT, in the hLHCGR and hFSHR, respectively). Thus, mutations in this motif influence receptor folding and PM localization.348,349 Mammalian cells expressing the recombinant hLHCGR display several distinct glycoprotein species with molecular masses (estimated from SDS gels) ranging from 65 to 240 kDa.296 The mature, PM-­expressed hLHCGR has been identified as an 85 to 95 kDa protein, whereas a 65 to 75 kDa band has been identified as an immature, partially glycosylated precursor that is located in the ER.296 The higher molecular weight bands ranging from 165 to 240 kDa are oligomers of the immature or mature receptors.296,463 Precursor, mature, and oligomeric forms of the hFSHR can also be detected in transfected cells but there is more variation in the reported molecular weights of these products. Estimates for the molecular mass of the mature cell-­ surface FSHR range from 74 to 80 kDa, whereas estimates for the molecular mass of the immature intracellular precursor range from 67 to 75 kDa.394,464,465 Higher molecular weight forms of the FSHR (∼170 kDa) have also been identified and appear to be oligomers of the immature intracellular precursor.465 It should be noted that many of the studies on the sizes and nature of the different forms of gonadotropin receptors have relied on immunological detection of epitope-­tagged receptors expressed in heterologous cell lines. Immunological detection of the endogenous gonadotropin receptors expressed in the testes or ovaries or in extragonadal tissues has been much more difficult because they are expressed at low densities and because of the nature of the gonadotropin receptor antibodies available. Using monoclonal or polyclonal antibodies (some of which have been rigorously validated), the mature and immature forms of the hLHCGR and hFSHR described above can indeed be immunologically detected in target tissues and homologous cell lines.296,466,467 Another posttranslational modification important for GPCR trafficking is palmitoylation. Cysteine residues in the carboxyl-­ terminus of several GPCR have been shown to be the target for S-­acylation with palmitic acid; in some receptors, this posttranslational modification is often required for efficient delivery of the protein to the cell membrane,468–472 where it facilitates anchoring of the receptor Ctail to the PM. The hFSHR exhibits in its Ctail two conserved cysteine residues (at positions 646 and 672) and one nonconserved cysteine residue at position 644. Although the hFSHR is palmitoylated at all cysteine residues, regardless of their location in the Ctail of the receptor,363 S-­acylation at C627 and C655 is not essential for efficient hFSHR PM localization, whereas at C629 it is, as replacement of this residue with glycine or alanine reduced detection of the mature form of the receptor by ∼40% to 70%. Further, when all palmitoylation sites are removed from the hFSHR, cell surface PM expression is reduced to ∼10% to 30% of that shown by the WT receptor.363 The hLHCGR is palmitoylated at two conserved cysteine residues (643 and 644),473 but in contrast to the hFSHR, palmitoylation of this receptor is

43

not important for trafficking to the PM, as the abrogation of palmitoylation did not appear to affect PM expression and agonist binding.362,473 The importance of palmitoylation of gonadotropin receptors in internalization and postendocytic processing of the PM-­expressed receptor following the formation of the hormone-­ receptor complex depends on the receptor. While in the hFSHR palmitoylation does not play any role in the internalization of the receptor/hormone complex, in the hLHCGR prevention of palmitoylation increased the rate of agonist-­stimulated internalization. In both receptors, abrogation of palmitoylation impairs receptor recycling to the PM and increases the fraction of receptor/hormone complex targeted to degradation via the proteasome/lysosome pathways (see below) (Fig. 2.11).361,362 Downward trafficking and the postendocytic fate of GPCRs, including the gonadotropin receptors, has received particular attention during the last two decades. It is currently considered a critical physiological event that determines not only the fate of the receptor but also an important second wave of signaling (Fig. 2.12).474–476 As previously mentioned, the number of active signaling receptors at the PM is classically regulated via mechanisms of receptor desensitization, endocytosis into the cell, and sorting to divergent intracellular fates (Figs. 2.11 and 2.12). Postendocytic sorting of internalized GPCRs is a complex process that regulates the fate of the receptor. Targeting GPCRs back to the PM (or recycling pathway) enables receptor recovery or resensitization, while alternatively the rapid delivery of GPCRs to the lysosome for degradation leads to permanent termination of receptor signaling.477,478 Most of the internalized rodents and hFSHR and hLHCGR recycle back to the cell surface. Both receptors exhibit particular residues and motifs that promote their recycling to the PM (Figs. 2.5 and 2.6).368,370,479,480 This is in contrast to rodent LHR, which are predominantly targeted to the degradative pathway because it lacks motifs present in the hLHCGR that promote recycling to the PM.480 In addition to palmitoylation and the GTALL motif present on the distal portion of the hLHCGR Ctail (Fig. 2.6), postendocytic targeting of this receptor is also influenced by an additional specific sequence in the Ctail of the receptor.368,370 Unlike the hFSHR, the hLHCGR exhibits a type 1 PDZ (for postsynaptic density 95/disc large/zonula occludens-­ 1)-­binding sequence or PDZ ligand in its C-­tail that binds the PDZ domain of the PDZ protein GIPC (Gαi-­interacting protein, C-­terminus), which functions as a scaffold to mediate receptor recycling from very early endosomes (VEE; see below) to the PM.479 In the case of the FSHR, recent data implicate APPL1 (adaptor protein containing PH domain, PTB domain, and Leucine zipper motif), which associates with the FSHR (see below) and that is present in a subpopulation of VEEs; this adaptor is able to bind GIPC and mediate receptor recycling.481 Further, PKA-­phosphorylated APPL1 (at Ser410) has been shown to be involved in receptor recycling and opposes cAMP signaling from VEEs (Fig. 2.12).475 However, the relationship between downward trafficking and receptor signaling is not limited to negative regulation of intracellular signaling from the PM; in fact, desensitization and internalization allow GPCRs to be differentially sorted to endosomes providing distinct intracellular membrane platforms either for activating or maintaining GPCR signaling, including heterotrimeric G protein signaling (Fig. 2.12), and to determine the fate of the internalized receptor (i.e., recycling vs. degradation). These endosomal intracellular signaling pathways exhibit distinct functions from those initially generated at the PM, demonstrating the complexity and integrated nature of trafficking and signaling.482 Instead of trafficking to the classically considered primary compartment for internalized GPCRs, the early endosomes (EE), both the hFSHR and hLHCGR, are rather targeted to the physically and functionally distinct VEEs, which lack EE and intermediate EE markers such as EEA1 (EE antigen), Rab5, and Fig. 2.12), where

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PART I  The Fundamentals of Reproduction Extracellular LH

FSH FSH

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Fig. 2.12 Models depicting the current understanding of LHCGR (left) and FSHR (right) endocytic pathways and the role of spatial control of receptor signaling.  Left: Following ligand binding, LHCGR activates a limited amount of Gαs-­mediated cAMP production from the plasma membrane. Through the recruitment of the ADP ribosylation factor nucleotide-­binding site opener (ARNO), and subsequent β-­arrestin (green polygon) release from membrane-­anchored small GTPase Arf6, the receptor engages β-­arrestin through its intracellular loop 3 at clathrin-­coated pits (CCP curved orange line leading to desensitization and internalization. While in CCPs, LHCGR also binds, through its Ctail, the adaptor protein GIPC (Gαi-­interacting protein, C-­terminus), which in turn drives LHCGR targeting to the very early endosome (VEE) where it dissociates. From APPL1-­positive VEEs, LHCGR is both able to recycle back to the plasma membrane and activate the majority of its Gαs/PKA/cAMP and ERK responses, potentially implicated in oocyte maturation and aromatase expression, respectively. Right: Upon FSH binding, membrane-­expressed FSHR activates G protein-­dependent cAMP production and acute pERK1/2 response. Activation of the receptor results in G protein receptor kinases (GRK) recruitment at the CCP and subsequent phosphorylation of FSHR Ctail, leading to receptor desensitization. GRK-­phosphorylated FSHR is recognized by β-­arrestins (green polygons), required for the sustained phase of FSHR signaling from the cell surface and internalization. Both the acute G protein-­dependent and the sustained β-­arrestin-­dependent signaling phases contribute to FSH-­induced mRNA translation. FSHR is then internalized and targeted to VEE in an APPL1-­promoted GIPC-­dependent manner. From APPL1-­positive VEEs, FSHR evokes cAMP and ERK signaling and recycling back to the plasma membrane. APPL1 positively regulates recycling and inhibits cAMP production.

they trigger cAMP and ERK signaling and recycle back to the PM (Fig. 2.11). As mentioned above, sorting of hLHCGR and hFSHR to VEEs is mediated via GIPC and promoted by APPL1, the latter also promoting recycling and inhibiting signaling (Fig. 2.12). This compartmentalized signaling appears to be functionally relevant for distinct functions of gonadotropin receptors such as regulation of oocyte maturation and steroidogenesis. Agonist stimulation of GPCRs is followed by a series of structural modifications and associations with scaffold proteins that eventually lead to effector uncoupling, internalization, and either recycling back to the PM or degradation in lysosomes and/or proteasomes. The hFSHR has been reported to be phosphorylated by second messenger-­dependent kinases PKA and PKC but also by GRKs 2, 3, 5, and 6 in various models.365–368,484 PKA and PKC contribute to both agonist-­dependent (homologous) and agonist-­ independent (heterologous) desensitization of the receptor. GRK-­ mediated phosphorylation leads to more complex effects as they are centrally involved in homologous desensitization while simultaneously regulating β-­ arrestin recruitment and subsequent β-­ arrestin effects on receptor internalization through clathrin-­coated pits and G protein-­independent signaling. As described above, a cluster of five serine and threonine residues located in the Ctail of the hFSHR (Fig. 2.5) has been shown to account for the bulk of

FSH-­induced phosphorylation as a result of GRK2 action.365 It has also been documented that β-­arrestins are recruited to the GRK-­ phosphorylated and agonist-­occupied FSHR.365–368,484 β-­arrestins recruited to GRK2-­or GRK5/6-­phosphorylated FSHR have been suggested to exert distinct intracellular functions365,485; GRK2-­ phosphorylated hFSHR predominates in the β-­arrestin-­mediated desensitization process whereas GRK5 and 6-­induced phosphorylation of the activated FSHR is required for β-­arrestin-­dependent signaling pathway in the immortalized cell line HEK293.365,485 It is well established that β-­arrestin 1 and 2 binding to GRK-­ phosphorylated FSHR leads to the internalization and recycling of the receptors.365,366,486 In contrast to the hFSHR, the hLHCGR does not recruit GRKs to promote its desensitization, which is instead mediated by the interaction of the receptor with ADP ribosylation factor nucleotide-­ binding site opener (ARNO), an exchange factor for ADP ribosylation factor 6 (ARF6), which then recruit β-­arrestins when bound to GTP.487 Here it is important to mention that recruitment to β-­arrestins bound by the hFSHR plays an important role in initiating distinct arrestin-­mediated signaling pathways, mainly the MAPK ERK, as will be described below. In the case of the hLHCGR, β-­arrestins are not apparently involved in MAPK-­ERK signaling, and thus this receptor does not demonstrate biased signaling through this particular signaling cascade.360

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

Extragonadal Expression LHCGR and the FSHR localize to gonadal cells, which contain their targets as evidenced by early hormone ablation experiments and subsequent development of in vitro bioassays. In the testes, the canonical location of LHCGR and FSHR is restricted to Leydig and Sertoli cells, respectively. In the ovary, expression of the LHCGR occurs in theca, interstitial, and granulosa cells from preovulatory follicles and in the corpus luteum, whereas the FSHR is limited to the granulosa cells of the maturing follicles. In all cases, one can easily demonstrate readouts following treatment with physiological levels of hormone and importantly the binding of a radiolabeled hormone to these tissues with high affinity and capacity. Certainly, the main physiological roles of the gonadotropin receptors can be attributed to their actions in the ovaries and the testes, as demonstrated by the phenotypes of individuals harboring activating or inactivating mutations of these genes (see below). Moreover, the phenotypes of genetically engineered mice with exaggerated or absent gonadotropin actions478,488–491 or mice with targeted inactivation or constitutively activated receptors343,492–495 appear to be explained entirely by the classical actions of LH and FSH in gonadal tissues. Nevertheless, in recent years there have been reports of low-­level FSHR expression in extragonadal tissues, where it has been proposed to have distinct physiological roles.496 The extragonadal sites where the FSHR has been detected include bone osteoclasts497,498 and monocytes,408,499 endothelial cells from umbilical vein,500 tumor vasculature and metastases,501,502 different sites of the female reproductive tract, the developing placenta,503 the endometrium,504 the liver,505 and the adipose tissue.506,507 Data on the expression (mRNA and protein) of either the FSHR WT or the splice variant originally identified in bovine granulosa cells402 in both osteoclasts and monocytes408 and, consequently, on the role of FSH in bone function are still quite controversial.497–499,508–512 Immunohistochemical studies employing monoclonal anti-­ FSHR antibodies have been the mainstay for documentation of the receptor because ligand binding is not detectable. The expression of the hFSHR in blood vessels from a number of malignant tumors501,513 and tumor metastases502 has been reported. Although the authors posit that FSHR at these locations may contribute to the growth and expansion of the tumor tissue by promoting angiogenesis, further studies are still necessary to validate these findings given that such claims can lead to unnecessary uncertainty in women undergoing hormone therapy for infertility or in menopause, when FSH levels are high. FSHR in several nonovarian reproductive tissues has been recently identified.500,503,504 Interestingly, fetoplacental haploinsufficiency of the Fshr in mice was associated with defects in placental growth and fetal loss in both heterozygous and homozygous null Fshr animals.503 More recently, a role for the FSHR in endometriotic lesions504 and in hepatic tissue505 has been proposed. The authors claimed that, at these locations, the FSHR might be involved in FSH-­ stimulated estrogen production and progression of endometriosis as well as in the expression of the low-­density-­lipoprotein receptor and regulation of low-­density-­lipoprotein cholesterol clearance, respectively. Nevertheless, further studies are still warranted to confirm the existence and elucidate more strongly the expression and physiological significance of extragonadal FSHRs and, if so, whether they are structurally similar to that primarily expressed in the gonads or are splice variants of the WT receptor that may be expressed in the PM in sufficient quantities to provoke a biological effect upon exposure to physiological concentrations of FSH. There is a large and equally controversial body of literature suggesting that functional LHCGR may also be expressed in a number of extragonadal tissues.514,515 The suggestion of extragonadal LHCGR expression has been based in many cases upon

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the detection of fragments of the LHCGR mRNA. Attempts to detect immunoreactive LHCGR protein often result in the identification of a protein (or proteins) that do not match the expected molecular weight of the authentic gonadal LHCGR (see above). A comparison of the actions of LH/CG in isolated cells or tissues from WT and LHCGR null mice would go a long way in shedding light on this controversy but curiously this has not been done often. A recent study employing different experimental approaches has identified potential angiogenic activity of hCG, which was abrogated by deletion of the LHCGR.516 This finding suggests that a functional LHCGR might be expressed in blood vessels. Further, the LHCGR has been detected in the secretory endometrium, particularly around the spiral arteries, as well as in the site of implantation in nonhuman primates.517 The authors claimed that the temporal expression of the LHCGR in this tissue might play an important role in inducing expression of specific genes important in modulating several processes, including decidualization, the immune system, cell survival, and the vascularization at the maternal-­fetal interface.517 Some of the clearest evidence for the extragonadal expression of a functional LHCGR comes from studies done on a woman who developed pregnancy-­associated Cushing’s syndrome and later developed it again after menopause.518,519 After menopause, there is clear evidence that the cortisol levels in this woman were controlled by LH because suppression of the hypothalamic/ pituitary axis with GnRH agonist controlled the hypercortisolism.518 Clinical findings in a number of other cases also seem to be explained by the inappropriate expression of the LHCGR in the adrenal cortex or in adrenocortical tumors.519,520 Ectopic expression of functional LHCGR in the adrenal cortex seems to be a common finding in transgenic or knock-­out mouse models with elevated levels of gonadotropins,520 and expression of the mature LHCGR can be readily documented in the adrenal glands of pregnant rats.521 Lastly, some reports indicate that during development other extragonadal rat tissues such as the kidneys express the immature form of the LHCGR, whereas nervous tissues express both the mature and immature forms.521,522 Direct action of gonadotropins on these tissues are still controversial.

Oligomerization A number of biochemical, pharmacological, and biophysical studies support the concept of receptor self-­association or oligomerization as a fundamental process enabling GPCR activity.523 It is currently accepted that multimerization of GPCRs is involved in fine-­tuning regulation of several physiological processes such as ligand binding, receptor trafficking, and regulation of PM expression levels as well as activation of signaling pathways.523,524 As with many other GPCRs, GPHRs also associate to form homodimers/oligomers and even heterodimers, in which two closely related receptors simultaneously expressed in the same cell associate (e.g., the LHCGR and FSHR in ovarian mature granulosa cells).6 In the case of the gonadotropin receptors and the TSHR, a number of studies have shown that these receptors self-­ associate.463,524–526 Although the structure of the FSH-­FSHRHB19 revealed that the FSHRED may form weakly associated dimers, with each molecule bearing one FSH molecule, later studies employing combined biochemical and biophysical approaches, directly demonstrated that the FSHR self-­associates early during receptor biosynthesis and that it can be identified as FSHR/ FSHR homodimers or FSHR/LHCGR heterodimers in the surface membrane of HEK293 cells.465,527 The mechanism and extent of FSHR self-­association are not known but it seems reasonable to assume that contacts occurring via the TMDs play an important role.462,528,529 The more recent crystal structure of the ligand-bound ectodomain of the FSHR20 demonstrated an additional mode of association of FSH with the entire FSHRED

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PART I  The Fundamentals of Reproduction

and a quaternary structure bearing three individual ectodomains, i.e., a trimeric receptor structure (Fig. 2.8A); this structure predicts hosting of one fully glycosylated FSH molecule or three deglycosylated molecules528 (Fig. 2.8B). In the former scenario, the FSHR may become activated, whereas in the latter the structure remains inactive, supporting early observations on the lack of effect of deglycosylated FSH for triggering signal transduction.530,531 The trimeric model also explains the reported differences in receptor binding activity between fully glycosylated and hypoglycosylated FSH isoforms, with the former exhibiting delayed and lower binding activities,532 and provides a mechanism for FSHR association with multiple effector proteins, thus drawing on several signaling pathways. The LHCGR also forms oligomers in the ER and at the PM, in a process that is unrelated to receptor activation.533 Importantly, coexpression of a misfolded LHCGR with its WT counterpart impairs the cell surface expression of the WT receptor and attenuates signaling,463,533,534 a dominant-­negative effect. Coexpression of splice variants of the LHCGR may also regulate the expression of the LHCGR and FSHR by forming intracellular oligomers that prevent the proper processing of the intracellular LHCGR precursor. For example, hLHCGR transcripts lacking exon 9 are prevalent in normal human ovaries but the resulting protein is not able to bind hCG or be adequately processed to be expressed at the cell surface.425,427 When coexpressed with the WT hLHCGR or hFSHR, the mutant lacking exon 9 associated with the immature forms of these receptors and exerted dominant-­negative effects by decreasing their cell surface expression.425,427 Formation of FSHR/FSHR complexes explains intermolecular complementarity between mutant FSHR dimers in which the function of a hormone binding-­deficient FSHR protomer may be rescued by a signal-­deficient counterpart following agonist exposure.535 Likewise, negative cooperativity can occur when upon ligand binding one protomer blocks or decreases the affinity of the second protomer for its ligand or intracellular signaling partners leading to a right shift of the dose-­response curve.536 Functional complementarity in the LHCGR has also been documented to occur both in vitro and in vivo.537–540 Complementarity between gonadotropin receptor homo-­or heterocomplexes conveys important physiological implications for the gonadotropin receptors as a temporal association and alternating negative cooperativity between the FSHR and the LHCGR in the ovarian granulosa cells. Complementarity between gonadotropin is evidenced by receptors homo-­or heterocomplexes. This observation of complementarity conveys important physiological implications for the gonadotropin receptors both in terms of temporal association and negative cooperativity between the FSHR and the LHCGR in the ovarian granulosa cells. This complementarity, might protect the follicle from premature luteinization or conversely from ovarian hyperstimulation, depending on the expression level of each receptor.541 Heterozygous mutations that deactivate receptors may remain silent by virtue of rescue with WT receptor or may conversely express as clinically adverse by virtue of the inactive receptor (i.e., with a trafficking defect), thereby preventing the native receptor from trafficking to the cell surface.462 A functionally important heterodimerization between the human FSHR and the G protein-­coupled estrogen receptor (GPER) has been recently characterized.542 In cultured human granulosa cells, FSHR-­ GPER heteromers are associated with reprogramming of FSH-­dependent cell death into proliferative stimuli, leading to oocyte survival. Specifically, no survival signals are detected at high FSHR:GPER ratios, which correlate with both coupling to the Gαs protein/cAMP signaling pathway and poor response to controlled ovarian stimulation in women undergoing assisted reproduction. In contrast, the presence of FSHR-GPER heteromers is preferentially associated with antiapoptotic/cell survival, pAKT-

via the Gβγ dimer, and a normal response to exogenous FSH administration.542 These data emphasize the importance of cross-­ talk between FSH-­and estrogen-­triggered signaling pathways in determining ovarian follicle selection in humans. In fact, previous data in mouse L cells stably expressing the human FSHR and the α-­ and β-­ estrogen receptors, demonstrated that FSH stimulation was associated with cell proliferation and transactivation of estrogen-­sensitive reporter genes.543 The occurrence of homo-­and heterodimerization as a mechanism for negative cooperativity or intermolecular complementarity between gonadotropin receptors as well as the recently reported heteromerization between the FSHR and GPER represents unique opportunities for therapeutic interventions.

Gonadotropin Receptor Signaling Pathways • In addition to well-­known canonical signaling cascades, additional pathways have been recently recognized, underpinning the complexity of signaling mediated by gonadotropin receptors. • Biased signaling in response to particular ligands and receptor modifications are now well-­recognized mechanisms by which gonadotropin receptors selectively activate particular intracellular signals.

Canonical Pathways, New Pathways, and Adapter Proteins Most investigators agree that the effects of the gonadotropins on the differentiated function of their target cells are mainly mediated by the activation of the canonical Gs/adenylyl cyclase/cAMP/PKA pathway. That pathway subsequently activates CREB, thereby modulating gene transcription.4,296,394,433,544,545 It is now clear that this is not the only signaling cascade activated by these receptors. Additional pathways are activated as detailed below and these may be involved in several gonadotropin-­dependent events such as proliferation and/or differentiation of target cells, and, at the molecular level, functional selectivity and differential gene expression. In recent years it has become evident that the FSHR is connected via conformational selectivity to a nonlinear and complex signaling network. It is a network mediated by several G protein subtypes, including the Gs, Gi, Gq/11, and Gh proteins,546–549 other types of receptors [e.g., the insulin-­ like growth factor 1 receptor and the epidermal growth factor (EGF) receptor (EGFR)], and the GPER.542,543,550 Other proteins associated with the receptor [e.g., β-­arrestins and Adapter protein-­containing Pleckstrin homology domain, Phosphotyrosine binding domain, and Leucine zipper motif (APPL)]346,551–553 further modulate those interactions. This conformational selectivity of FSHR interactions promotes activation of a number of signaling pathways including those mediated by distinct kinases (such as PKA, PKC, PI3K, PKB/Akt, and ERK1/2) including signaling in VEEs.297,365,475,547,548,550,554–556 That complex network of pathways allows for fine-­tuning regulation of the gonadotropic stimulus, where the activation/inhibition of its multiple components may vary depending on the cell context, developmental stage of the host cells (Sertoli and ovarian granulosa cells), and concentration of receptors and ligands.321,550,555,557,558 Of particular interest are FSHR-­mediated pathways triggered by cAMP accumulation but independent from PKA activation, such as those activated by the exchange protein directly activated by cAMP (EPAC). In addition to CREB, cAMP-­activated PKA activates p38 MAPKs, ERK 1/2, p70S6 kinase (p70S6K), and PI3K via insulin receptor substrate 1 (IRS-­1), leading to PKB/Akt and FOXO1 transactivation and thereby to activation/repression of FSH-­regulated genes.559­561 Meanwhile, EPAC promotes p38 MAPK and PKB/Akt phosphorylation, and upregulation of the EGFR.554,562 In addition to G proteins, FSHR also associates with other cytoplasmic interacting proteins including the APPL1,2 and 14-­3-­3τ adapters and the scaffolds β arrestin 1,2.345 346 365 552 563 Linking of the FSHR with APPL1 occurs at the IL1 of the receptor, specifically at K393553

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

(Fig. 2.5), whereas the association with 14-­3-­3τ maps to the IL2 and overlaps with the canonical G protein binding sites.345,346 Agonist stimulation rapidly induces FOXO1a phosphorylation in HEK293 cells leading to inhibition of apoptosis via either serum and glucocorticoid-­induced kinase (Sgk) or APPL1 interaction with the upstream PI3K/Akt pathway.554,563,564 In addition, APPL1 also participates in the FSH-­induced inositol 1,4,5 triphosphate (IP3) pathway and implicates it in intracellular calcium signaling in granulosa cells.553 The coexistence of APPL1 and 14-­ 3-­3τ associated with the FSHR, suggests that FSH causes phosphorylated FOXO1a to be sequestered by 14-­3-­3τ with APPL1 facilitating this antiapoptotic process.346 The FSHR activates ERK1/2 phosphorylation and signaling, which have been shown to play an important role in immature granulosa cells on the induction of a subset of FSH gene targets that define the mature preovulatory granulosa cells, including the genes encoding the α-­subunit of inhibin, the LHCGR, the EGFR, and the enzyme aromatase.565,566 ERK1/2 activation occurs through two distinct pathways, one mediated by Gsα and the other by β-­arres tins,365,567,568 which are primarily involved in agonist-­ stimulated desensitization and internalization of the FSH/FSHR complex (see above). As discussed previously, phosphorylation by GRK5 and 6 is required for ß-­ arrestin-­ dependent, Gsα-­independent signaling.365,485 When activated by the Gsα-­PKA pathway, ERK1/2 phosphorylation occurs via PKA-­promoted dissociation of ERK from a 100-­kDa phosphotyrosine phosphatase that inhibits ERK activation and its translocation to the nucleus.568 PKA-­stimulated ERK activation occurs early and is transient, peaking at 5 to 10 min after FSH exposure, whereas in the Gsα-­independent pathway mediated by β-­ arrestins, activation is slower and sustained and mainly occurs during the ensuing 10 to 30 min of agonist exposure.365 LH is essential to stimulate androgen production by the Leydig and theca cells as well as for ending the FSH-­regulated program for steroidogenesis and granulosa cells growth, and for promoting the luteinization of cells into luteal cells.555 LH also induces genes necessary to promote oocyte maturation, follicle rupture, and ovulation. To do this, the LH-­activated LHCGR must turn on a variety of signaling pathways. The LHCGR was one of the first GPCRs shown to independently activate distinct G protein-­dependent signaling pathways, adenylyl cyclase (through Gsα) and phospholipase C (through Gαq/11).569,570 Although this observation was initially made in heterologous cells expressing the recombinant mouse LHCGR, it has now been extensively reproduced in a variety of cell lines transfected with either the rodent or human LHCGR.296 LHCGR-­ mediated activation of the Gsα/PKA/cAMP pathway leads to activation of several downstream signaling cascades, such as the EGF network, which includes expression of the EGF-­like growth factors amphiregulin (AREG), epiregulin (EREG), and betacellulin (BTC) that bind and transactivate the EGFR in the mural and cumulus cells in an autocrine/paracrine fashion, thus leading to activation of the ERK1/2 cascade and the expression of genes necessary for cumulus expansion.571–577 Activation of the EGFR-­regulated network is also one important factor involved in the resumption of oocyte meiosis as it promotes the decline in cGMP levels, whose concentrations negatively regulate the activity of the phosphodiesterases necessary to reduce intraoocyte cAMP and promote meiotic resumption.572–574,576,578-­582 The ERK1/2 cascade, activated through EGFR-­Ras and also probably through cAMP and/or PKC, is critical for several LH-­stimulated processes (including androgen production)583,584 and those occurring around ovulation (such as cumulus expansion, oocyte resumption of meiosis, ovulation, and luteinization).555,574,585,586 Further, it has been shown that most of the ovarian genes that are up-­or downregulated by LH are targets of ERK1/2.555,574 An additional level of regulation controlled by the ligand-­ activated FSHR occurs at the level of mRNA translation through a cross-talk of PKA with Akt-dependent pathways to stimulate mammalian target of rapamycin (mTOR) and P70S6

47

kinase558,587,588 leading to mRNA translation and regulation of cell cycle progression and cell proliferation.544 It is thought that this FSH-­dependent regulation of mRNA translation stimulates granulosa cell proliferation via Gαs-­dependent ERK-­mediated phosphorylation of the mTOR effector, TSC2 (tuberin).589 Activation of tuberin stimulates P70S6K activity resulting in increased enhanced cyclin D2 expression.544 Further, FSH activation of the mTOR pathway leads to the expression of follicular differentiation markers, including the LHCGR, inhibin-­α, aromatase, and βII subunit of PKA. These observations emphasize the importance of the agonist-­ activated FSHR signaling network in regulating the physiological functions of gonadotropin. Moreover, the effects of FSH signaling on mRNA transcription and translation are also extended to regulation of microRNAs (miRNA) that play an important role in reproductive function. As miRNAs may destabilize particular mRNAs and regulate translation, they play an important role in controlling the FSH signaling network and expression of genes involved in meiosis and spermiogenesis.590–592 A similar situation occurs in the ovarian follicle, where expression of a number of miRNAs [e.g., miR10] are regulated by FSH, impacting follicular growth, progesterone, estradiol synthesis, and/or LHCGR expression.435,593–595 Studies in several experimental models have found that LHCGR activates members of other families of G proteins, including the Gαq/11 protein,570,596–598 which contributes to the ovulatory process via the progesterone receptor as has been demonstrated in mice with a granulosa cell-­specific deletion of Gαq/11.599 Although these Gαq/11-­deficient mice resumed meiosis and showed cumulus expansion upon hCG exposure, they failed to increase inositol phosphate accumulation and express the progesterone receptor and exhibited marked subfertility because of trapping of the oocytes destined for ovulation in preovulatory follicles or corpora lutea.599 LHCG-­activated Gαq/11 and phospholipase C is also involved in Ca2+ mobilization from intracellular stores in a process potentiated by receptor cross-­ talk through heterodimerization with the unliganded hFSHR. That association promotes a conformational change in the hLHCGR, thereby presumably favoring on the one hand LH binding and LH/hLCGR-­mediated Gαq/11-­βγCa2+ signaling, and on the other hand inhibiting hFSHR-­dependent cAMP production.537,541 Although Ca2+ is not necessary for oocyte maturation, it is apparently involved in luteinization.600 In addition, LHCGR activates PI3K and PKB/Akt in a cAMP-­independent fashion.571,572 It is thought that this pathway is involved in oocyte meiotic resumption, albeit in a dispensable manner given that mice lacking PTEN (phosphatase and tensin homolog deleted on chromosome 10, which functions as a negative regulator of PI3K action) or PDK (phosphoinositide-­ dependent kinase, which phosphorylates PKB/Akt) did not exhibit alterations in meiosis II progression or germinal vesicle breakdown.601–603 The LHCGR also activates the mammalian target of rapamycin complex 1 (mTORC1) signaling through the cAMP/PI3K/Akt signaling pathway to regulate cell proliferation and expression of several key enzymes involved in androgen biosynthesis, including the P450 side-­chain cleavage enzyme (P450scc), 3β-­hydroxysteroid dehydrogenase type 1, and 17α-­ hydroxylase/17,20lyase (P450c17), without modifying the expression of the steroidogenic acute regulatory protein (StAR) in theca-­interstitial cells.604,605 Although the LHCGR recognizes both LH and hCG ligands, in vitro studies on different cell lines expressing this receptor (e.g., COS-­ 7, HEK-­ 293, granulosa-­ lutein, and mouse tumor Leydig cells) have suggested that both gonadotropins are distinguished by the LHCR (probably at the level of exon 10) and differentially activate downstream signaling pathways (e.g., cAMP production and MAPK phosphorylation) both quantitatively and qualitatively. Thus, with distinct kinetics and time frames of gonadotropin exposure, hCG is more potent in stimulating

2

48

PART I  The Fundamentals of Reproduction

cAMP production and steroidogenesis and LH is more active to induce ERK1/2 and AKT activation.321,606,607 Interestingly, the agonist effect of LH on progesterone production and β-­arrestin 2 recruitment in mouse tumor Leydig cells was rather partial compared to that evoked by hCG, but not on testosterone biosynthesis where the effect was similar for both gonadotropins.607 These findings implicate β-­arrestins in the regulation of steroidogenesis and further emphasize the differential effects of these hormones depending on the desired biological effect and primary cell context (i.e., progesterone production by the corpus luteum during early pregnancy vs. testosterone production at the Leydig and/ or theca cells). That the LHCGR has the ability to discriminate between the two ligands and evoke biased signaling might have pharmacologic implications in the clinical arena, specifically with regard to their use in assisted reproduction and treatment of hypogonadism. In summary, a number of studies in experimental animal models and in humans indicate that several gonadotropin-­regulated intracellular signaling pathways must coalesce to coordinately regulate distinctly different cellular processes involved in steroidogenesis, follicular growth and maturation, cell differentiation, and ovulation. These signaling cascades are activated in a time-­regulated fashion depending on the differential expression of gonadotropin receptors during the different stages of the ovarian cycle. Thus, granulosa cells from small antral follicles express the FSHR, with a progressive increase during follicular maturation. LHCGR expression initially confined to the theca-­ interstitial cells is also induced to express in antral follicles, and expression increases during follicular growth in response to FSH, FSH-­stimulated estrogen production, and other paracrine factors. In fact, both gonadotropin receptors are simultaneously expressed in granulosa cells from mature, preovulatory follicles, in which activation of the LHCGR by its cognate ligand eventually suppresses the effects of FSH on steroidogenesis and follicle growth and promotes the ovulatory process. After ovulation and differentiation of granulosa cells into luteal cells, LHCGR expression regulates progesterone and estradiol production by the luteal cells until corpus luteum regression occurs.555,608,609

Biased Agonism It is currently accepted that preferential activation of distinct GPCR-­mediated signaling cascades may occur through biased signaling that results from the stabilization of distinct receptor conformations353,610,611 in response to particular ligands or caused by receptor mutations.611–614 In the case of the FSHR, functional selectivity or biased agonism towards Gi-­ mediated signaling has been documented for some naturally occurring glycosylated variants of human FSH615–617 and recombinant FSH expressed in insect cells618 as well as for selective allosteric compounds with FSH agonist or antagonist properties605,619,615,616 Bias towards β-­arrestin-­mediated ERK1/2 MAPK signaling also has been observed when FSHR is treated with eCG/anti-­eCG antibody complexes generated from a large number of eCG-­ treated goats620 as well as with a modified agonist [truncated equine LHβ (Δ121–149) combined with Asn56-­deglycosylated equine LHβ].621 This latter molecule is particularly interesting because while preferentially activating the β-­arrestin-­ ERK1/2 phosphorylation module, it concomitantly behaves as an antagonist of FSH-­ stimulated cAMP-­ PKA signaling when tested at equimolar concentrations.621,622 Similarly, in HEK293 cells expressing FSHR, hypoglycosylated FSH18/21 exhibited a preference towards β-­arrestin-­mediated ERK1/2 activation when compared viz à vis with its fully glycosylated counterpart (FSH24). Additionally, the mechanism whereby hypoglycosylated FSH18/21 promoted intracellular Ca2+ (iCa2+) accumulation also differed from FSH24 in that iCa2+ mobilization stimulated by the former depended more on release from intracellular stores than

on influx from Ca2+ channels, indicating differences in mechanisms through which glycoforms promote iCa2+ accumulation.33 At the receptor level, biased agonism has been documented when hFSHR PM expression is severely impaired as a result of a loss-­ of-­function mutation (A189V, see below)623 (Fig. 2.5); in this case, β-­arrestins recruited to the agonist-­bound receptor assembled a MAPK module, whereas G protein-­dependent signaling remained impaired.624 Also interesting are the missense M512I and I423T FSHR mutations, which led to an impaired FSH response in terms of cAMP accumulation and PI3K activation but not ERK1/2 phosphorylation.625,626 Also, swapping K589 and A590 in the EL3 of the hFSHR with their respective residues in the hLH/CGR (A589N and A590S hFSHR mutants) led to an increased rate of internalization of the receptor and preferential signaling through the cAMP pathway over the MAPK-­ERK 1/2 one, indicating that those residues in the EL3 are important in regulating the latter signaling cascade at the postendocytic level.331 Conditional signaling (biased signaling dependent on the cell context) at the LHCGR has been recently reported for two negative allosteric modulators of the FSHR (see below),619 in which they differentially inhibited CG-­mediated steroidogenesis when tested in the mLTC-­1 cell (a mouse Leydig cell tumor-­ derived cell line) and rat primary Leydig cells.627 Although the physiological role of biased signaling at the gonadotropin receptors remains to be defined, this phenomenon opens the door for the design of drugs that may positively or negatively regulate selective signaling pathways on the follicle and that may be potentially useful in the clinical arena to treat estrogen-­and/or androgen-­dependent diseases and infertility as well as in contraception.

Gonadotropin Receptor Clinical Significance • Naturally occurring mutations in the gonadotropin receptor genes, albeit rare, lead to human disease. • Mutations in the gonadotropin receptor genes may cause loss or gain of function of the altered receptor depending on the particular location and nature of the substitution at the receptor protein. • Polymorphisms in receptor genes may necessitate personalized therapeutic intervention and inform human genetics.

Naturally Occurring Mutations A number of naturally occurring mutations of the LHCGR and FSHR associated with human reproductive disorders have been reported. These are shown in Figs. 2.5, 2.6, 2.13, and 2.14 and have been extensively reviewed.133,297,343,628-­632 Mutations in gonadotropin receptors may lead to either loss-­of-­function or gain-­of-­function of the receptor protein, depending on the location, nature of the amino acid substitution, and how the mutation alters receptor conformation and function. Loss-­ of-­ function mutations in the gonadotropin receptor genes may lead to disease, whenever both alleles are affected by a mutation, as it occurs in individuals who are homozygous or compound heterozygous for mutations in the hFSHR or hLHCGR genes. Several naturally occurring inactivating mutations scattered throughout the polypeptide sequence of the gonadotropin receptor molecules have been described. In 46, XY individuals, loss-­of-­function mutations in the hLHCGR gene lead to different phenotypes, including severe genital ambiguity [Type 1 Leydig cell hypoplasia (LCH)]. This is due to the effect of the mutation on the fetal Leydig cell response to the hCG stimulus, which is a determinant for male sexual phenotypic differentiation. It also leads to cryptorchidism and micropenis (Type II LCH). Although women with inactivating mutations in the hLHCGR display pubertal development, they frequently exhibit primary or secondary amenorrhea and infertility, or failure to respond to controlled ovarian hyperstimulation due to the development of

49

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

A

2

B

N-terminus

N-terminus

G117R V144F Q170 Stop

I152T

EL1

EL3

I114F

Extracellular EL2

C131R

/ CG

F194V

H1 Q303 Stop

Y331

Hinge S293 Stop Q246 Stop C343S R283 Stop Q525 Stop EL3 A593P I528 Stop E354K Extracellular A589 Stop L608 S586 Stop I585 Stop deIL608-609 H6 Y612 Stop C543R C545 Stop W465 Stop

Membrane R554 Stop

H3

H5

Leucine-rich repeat domain

C581R L368P

D578F/Y/ E/H/A

EL1

A373V

I457R

EL2

L502P

H7

I542L

IL3

C617Y

H2 H1

I415T A449T S616Y C617 Stop W491 Stop

N400S I374T N377D

H7

T577I

I575L

I625k

Membrane

M398T

Intracellular

A572V

H8 M571I

R479 Stop

H6

H5

Intracellular

H2

A568V

C-terminus

T3921

H8

IL1 IL3

D564G

C-terminus

Fig. 2.13 Positions of naturally occurring mutations at the LHCGR in a homology model complexed with the heterodimeric human CG crystal structure.  Aside from X-­crystallographic structures of the hormone CG, for the LHCGR, no direct structural information is yet available. However, because of high sequence similarities between FSHR and LHCGR as well as between FSH and hCG, general structural properties of the already determined FSHR LRRD/hinge-­FSH complex solved by X-­ray crystallography (Fig. 2.7) can also be assumed—in approximation—for a homology LHCGR/CG complex model. The receptor (beige backbone cartoon) likely binds the hormone (hCG subunits α—magenta, β—blue, surface presentation) extracellularly between the leucine-­rich repeat domain (LRRD) and the hinge region (brown). Additionally, a sulfated tyrosine (cyan) at the hinge region interacts with the hormone CG, also known from the FSHR/FSH complex (Figs. 2.5 and 2.14). In principle, the LRRD, the hinge region, and the serpentine domain are arranged sandwich-­like, whereby several disulfide bridges between the LRRD and the hinge region attach these fragments tightly together. The intra-­(IL) and extracellular loops (EL) connect the transmembrane helices (H) 1 to 7. The internal agonist sequence 350FNPCEDIMGY359 in the hinge-­region/helix 1 transition is colored magenta. In this model positions of naturally occurring inactivating mutations (red atom-­ spheres) (A) and constitutively activating single point mutations (CAMs, green atom-­spheres) (B) are mapped. This 3-­dimensional view highlights that CAMs are located mostly in the center of the transmembrane core at helix 6. In contrast, inactivating mutations occur across the entire receptor structure (A). The inactivating mutations can modify the capability of the receptor to interact with the hormone (mutations located in the LRRD or hinge region), activate the receptor in the presence of ligand, interfere with the activation of the coupled G-­protein(s) (located intracellularly), or lead to a misfolded receptor protein and subsequent intracellular retention. (The figure was kindly provided by Gunnar Kleinau, Charité-­Universitätsmedizin Berlin, Berlin, Germany).

genuine empty follicle syndrome.633,634 Meanwhile, inactivating, loss-­of-­function mutations of the hFSHR in men lead to impaired quality of spermatogenesis with normal testosterone production, which probably contributes to fertility preservation.635 In women with inactivating hFSHR mutations, the panorama is completely different and comprises an array of phenotypes, ranging from lack of pubertal development and primary amenorrhea with the arrest of follicular maturation between primordial and preantral stage and complete resistance to FSH stimulation to secondary amenorrhea and premature ovarian failure.623,636,637 In either case, the level of residual, functional receptors at the PM has been shown to correlate with the severity of the clinical phenotype expressed by the patients bearing inactivating mutations in these receptors628-

response to exogenous gonadotropins and other potentially useful drugs, such as receptor allosteric modulators (see below). Inactivating mutations of the gonadotropin receptors are germline, missense, or nonsense mutations that result in single amino acid substitutions in the receptor protein (Figs. 2.5, 2.6, 2.13A, 2.14) or introduction of a stop codon in their corresponding mRNAs, leading to amino acid deletions or insertions, or premature truncations of the receptor. Due to the scattered distribution of the mutations along the sequence of the receptor, the mutations may compromise the synthesis of the receptor protein due to large truncations (Class I GPCR mutations452), domains involved in agonist binding (Class III mutations), or receptor activation or coupling to effectors (Class IV mutations). Frequently, the mutations also lead to misfolded, trafficking defective proteins

50

PART I  The Fundamentals of Reproduction

TABLE 2.1  Functionally Characterized, Trafficking Defective hLHCGR Mutations

I61N

N-terminus

K140 Stop R59 Stop

Leucine-rich repeat domain N191I

S128Y

/ FSH

G216R I160T

Y335

A189V

V221G

D224V

M265V

M512I

Hinge

T438I V514A A433D

P348R P519T L601V

EL2

EL3

L597I P504S

H2

Extracellular

T449A,I,N

I423T

F591S

I418S

P587H I545T

N431I

EL1

H3

A419T I411N

H5

H6

H7

H1

A575V

D408Y

IL1

Membrane

H8

Intracellular

R573C

D567G,N

IL3

R634H

C-terminus

Fig. 2.14 Positions of naturally occurring mutations at the FSHR-­ FSH complex model.  Structures of the extracellular FSHR complex constituted by the hinge region (brown), the leucine-­rich repeat domain (LRRD), and heterodimeric human FSH (FSHβ (orange) and FSHα (magenta), surface representation) have been determined previously.20 The hinge region links the LRRD with the 7-transmembrane domain (7TMD) of the FSHR (white-­gray backbone cartoon), which comprises seven transmembrane helices (H1–H7) interconnected by intracellular (IL) and extracellular loops (EL). The 7TMD was modeled based on the known active-­state structure of the β2-­adrenergic receptor/Gs complex.352 The exact orientation between the FSH/LRRD/hinge region complex and the 7TMD is still unclear. However, the spatial localization of naturally occurring mutations can be studied at this model assembly and is highlighted by the side chains of wild-­type positions (atom spheres, without hydrogens). Loss-­of-­function mutations are indicated by red color and gain-­of-­function mutations by green color. Gain-­of-­function mutations are characterized by an increase in ligand-­independent basal signaling activity or by causing promiscuous (e.g., hCG) ligand binding (lilac spheres and labels). The V514 mutant (magenta circle in EL2) led to increased plasma membrane expression and OHSS at low FSH doses. The internal agonist sequence 353FNPCEDIMGY362 in the hinge-­region/helix 1 transition is colored magenta. (The figure was kindly provided by Gunnar Kleinau, Charité-­ Universitätsmedizin Berlin, Berlin, Germany.)

unable to transport from the ER to the PM (Class II mutations) (Fig. 2.11). These functional defects are not mutually exclusive, as one mutation may lead to functional defects in both intracellular traffic and any other function. This is the case, for example, of the delLeu608/Val609 misfolded hLHCGR mutant, which in spite of being expressed at low levels at the PM and exhibit ing normal binding affinity, is unable to activate the G

Region

Mutations

Extracellular domain or I114F, Val144Phe, hinge region Phe194Val, del exon 8, del exon 10, R283 STOP, Cys343Ser, In-­frame insertions Gln18_Leu19ins11 and immediately Gln18_Leu19ins9 upstream of the predicted signal peptide cleavage site Signal peptide Leu10Pro transmembrane [Thr392Ile, Ile374Thr + domains (TMDs) Thr392Ile, Ile415Thr, Thr461Ile, Leu502Pro, Cys543Arg, Ala593Pro, delLeu608-­Val609, Ser616Tyr, C617Lfs STOP, and Ile625Lys

References 632, 639, 645–651 645, 649, 652

650 638–640, 647, 649, 652–654

upon exposure to the agonist.638 The same appears to occur with the Ile625Lys hLHCGR mutant, which leads to partial LCH in which in vitro studies showed that in addition to low PM expression, the mutant did not couple with effectors efficiently.639,640 Since in addition to provoking defective folding and intracellular retention of the protein, the mutations may also affect an intrinsic function of the receptor (e.g., binding to agonist or signal transduction), it might be expected that the benefit of treatment with pharmacological chaperones (or pharmacoperones) to correct folding and trafficking will be limited. Nevertheless, the possibility exists that the pharmacoperone may correct not only trafficking of the receptor to the PM but also receptor function by virtue of modifying the conformation affected by the mutation that primarily impacts ligand binding or signal transduction. This effect has been reported for the cell-­permeant, allosteric small molecule agonist Org 42599/Org 43553, which corrected function of the binding deficient Cys131Arg and Ile152Thr or the Glu354Thr signaling deficient hLHCGRs.641 Among the 40 or so inactivating hLHCGR mutants described so far (Figs. 2.6 and 2.13A), ∼40% are nonsense or frameshift mutations and at least ∼50% are trafficking defective receptors in which the net amount of functional receptors expressed at the PM is decreased to a variable extent.633,642–644,632 The phenotype of individuals harboring these types of inactivating mutations of the hLHCGR is, therefore, defined by the density of residual and functional receptors (not retained intracellularly) that reach the PM and bind agonist. These functionally characterized trafficking-­defective hLHCGR receptors bear mutations either in their ECD or hinge region632,639,645–651 or the TMDs 638–640,647,649,652–654 (Table 2.1). Some of these mutant hLHCGRs are interesting from structural and functional points of view. The Phe194Val misfolded mutant bears the substitution in a highly conserved motif of the gonadotropin receptors (193Ala-­Phe-­Asn-­Gly-­Thr197 at the hLHCGR ectodomain) that contains the Asn-­Gly-­Thr glycosylation motif. This mutation severely impairs the trafficking of the mutant receptor to the PM without altering agonist affinity.646 In the case of the inactivating Ala593Pro and Ser616Tyr mutant hLHCGRs, which leads to severe or moderate forms of LCH, respectively, both exhibited normal ligand binding affinity but the response to agonist was absent or severely impaired due to misfolding and intracellular retention of the majority of mutant receptor synthesized.458,647 Further, it was shown that these particular mutants have distinct conformations and display different folding conformations during their maturation at the ER.458 Interestingly, Newton and colleagues655 found that the Org 42599/Org 43553 compound partially rescued folding, PM

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

expression, and the function of these two mutants. The 1850delG mutation in exon 11 is also interesting; this is a frameshift mutation that results in replacement of the last 83 amino acid residues of the receptor by a 21 amino acid sequence lacking the traffic-­ regulating Phe(X)6Leu-­Leu motif at the amine-­terminal end of the Ctail (see above). Since this mutant is not a misfolded protein but rather a mutant lacking an important sequence critical for intracellular traffic of the receptor, its functionality could not be rescued by pharmacological chaperones.656 The study of the molecular physiopathogenesis of other mutations has unveiled important information on the structure-­ functional relationships of the LHCGR. Mutations in exon 6A lead to LCH Type 1 or 2 (A557C,G558C and 580A>C). If the last two mutations occur in a compound heterozygous state with the Thr461Ile and the Ile415Thr mutations, respectively, it will provoke an aberrant nonsense-­mediated mRNA decay of the internal exon 6A transcripts. This will lead to altered ratios of LHCGR transcripts that yield predominantly nonfunctional LHCGR variants or translation products of the abnormal transcript that inhibit function of the normal receptor through a dominant-­negative effect.428,429 The phenotype associated with the deletion of exon 10 also deserves special mention. This mutant was initially found in a 46XY individual who failed to undergo puberty in spite of normal sex differentiation.657 When expressed in a heterologous cell line, an hLHCGR construct lacking the amino acids encoded by exon 10 (residues 290–316 in the hinge region of the ectodomain) (Fig. 2.6) was transported properly to the cell surface, bound both hLH and hCG with high affinity, but its sensitivity to hLH (and not to hCG) was reduced about 40-­fold.658 Therefore, the residues encoded by exon 10 of the hLHCGR appear to be involved in receptor activation by hLH but not by hCG. In this vein, the CGR of the marmoset monkey provides an interesting evolutionary parallel to the mutant hLHCGR variant lacking exon 10. Since the marmoset pituitary does not express LHβ659 and exon 10 of this monkey LHCGR gene is spliced out of the mature mRNA, this truncated receptor only binds and responds to hCG.660 Gain-­of-­function mutations in the LHCGR lead to constitutively active mutants (CAMs) (Figs. 2.6, 2.13B, and Table 2.2). Numerous hLHCGR CAMs have been reported, not the least because of the dramatic phenotype of familial gonadotropin-­ independent isosexual male-­ limited precocious puberty.132 In early studies, this rare disease was initially named “familial testotoxicosis” to emphasize the autonomous nature of testosterone hypersecretion present in affected subjects.132,661,662 The cardinal biochemical features in affected boys are elevated levels of circulating testosterone in the face of undetectable serum immunoreactive levels of LH.663,664 Signs of puberty appear early in childhood, by 2 to 6 years of age, and the testicular histology is characterized by Leydig cell hyperplasia.661,662,664 The pattern of inheritance is autosomal dominant, albeit sporadic cases have also been reported.665 The phenotypic expression is variable and also may show incomplete penetrance, as exemplified by the cases with the Met398Thr mutation, in which some male family members who are carriers of the mutation exhibit precocious puberty phenotypes.666 This suggests that other factors, such as interaction with a modifier gene that silences the activating effect of the amino acid change, may impact the expression of the clinical phenotype. A still remaining enigmatic situation is the apparent lack of phenotype in women with hLHCGR CAMs. In this vein, the phenotypic features of an animal model bearing an LHCGR CAM have been recently reported; knock-­in female mice expressing the Asp582Gly (Asp578Gly in human) LHCGR CAM exhibited precocious puberty, infertility, irregular estrous cyclicity, elevated steroid hormone levels, polycystic ovaries, and granulosa cell tumors.667 These contrasting phenotypes (i.e., normal phenotype in women versus a highly altered phenotype in female mice) may be due to several factors, including species differences

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TABLE 2.2  Naturally Occurring CAMs of the LHCGR and FSHR, and the Corresponding Mutations at Homologous Positions in the TSHR (Reproduced from Ulloa-­Aguirre A, Reiter E, Bousfield G, Dias JA, Huhtaniemi I. Constitutive activity in gonadotropin receptors. Adv Pharmacol. 2014;70:37–80. with permission from Elsevier Inc [2014]) Location LHCGR TMD1 TMD2 ECL2 TMD3 TMD5 TMD6

TMD7

Leu368(1.41)Pro*

TSHR

FSHR

-­ -­ -­ Asn431Ile Leu457(3.43)Arg Leu512Arg/Gln -­ Thr449(3.32)/Ile/Ala Ile542(5.54)Leu Val597Phe/Leu Ile545Thr Asp564(6.30)Gly Asp619Gly Asp567Gly/Asn Ala568(6.34)Val Ala623Ile/Ser/Val -­ Met571(6.37)Ile Met626Ile -­ Ala572(6.38)Val Ala627Val -­ Ile575(6.41)Leu Ile630Leu/Met -­ Thr577(6.43)Ile Thr632Ala/Ile -­ Asp578(6.44)Gly/ Asp633Ala/Glu/His/Tyr -­ Tyr/Glu/His Cys636Trp -­ Cys581(6.47)Arg Cys617(7.47)Tyr Cys672Thr -­ Ala373(1.46)Val Met398(2.43)Thr

-­ Ala428Val Met453Thr

*(Generic amino acid numbering according to the Ballesteros and Weinstein nomenclature [354].)

in the regulation of the LHCG expression and function and the fact that in women the basal activity of the receptor in the absence of ligand is modest. It is much higher in mice. There is a 3-­to 5-­fold increase in basal cAMP compared with the WT receptor in women versus a 23-­fold difference in mice).668 To date, 14 naturally occurring gain-­of-­function mutations in the hLHCGR have been reported.343 In contrast to the scattered location of the naturally occurring loss-­of-­function mutations of the hLHCGR, all naturally occurring CAMs reported to date are localized in exon 11, which codes for the carboxyl-­terminal end of the hinge region of the ECD and the 7TMD and intracellular regions of the hLHCGR (Figs. 2.6 and 2.13B).343,631 More specifically, hLHCGR CAMs arise from amino acid substitutions in the serpentine domain of the receptor protein, particularly in the α-­ helices that conform to the TMDs or in the junction of the helices with the intracellular loops (e.g., the Asp564Gly mutation) (Figs. 2.6, 2.13B and Table 2.2). The first LHCGR CAM identified was located at amino acid 578 (Asp578Gly) at the TMD6,669,670 and 7 additional locations in this helix have since then been identified as sites for CAMs of this receptor.663,664,669–675 Thus, helix 6, and particularly Asp578, are hot spots for naturally occurring hLHCGR CAMs. The remaining LHCGR CAMs are located in the remaining TMDs, with the exception of TM4.663, 666, 671, 674–677 Several major features characterize LHCGR CAMs; these include:    A.  They are located at highly conserved residues among the GPHRs. In fact, TSHR CAMs at these very same conserved residues have also been identified in patients with autonomous thyroid adenomas (in the case of somatic mutations) or hereditary/sporadic nonautoimmune hyperthyroidism (germline mutations) (Table 2.2).678 This suggests that these receptors share common mechanisms of activation. As in the case of the hLHCGR CAMs, the TMD6 of the hTSHR is a hot spot for naturally occurring mutations.678 In contrast, only two of the four hFSHR CAMs identified to date (Ile545Thr at the TMD3 and Asp567 at the cytoplasmic face of TMD6) share locations with the LHCGR and TSHR CAMs.679–681 B.  When expressed in heterologous cell types, hLHCGR CAMs display variable levels of constitutive (i.e., hormone-­ independent) activity. The addition of hLH or hCG to cells expressing these mutants may or may not result in additional

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PART I  The Fundamentals of Reproduction

activation.631 That probably reflects both stabilization of the receptor in intermediate states of activation and secondarily the strong, cAMP-­mediated stimulation of phosphodiesterase activity in the basal state.677,682 On the other hand, it has been shown that different CAMs are able to stimulate multiple G proteins and second messengers in the basal state. This is the case, for example, of the Leu457Arg and Asp578Tyr CAMs, which activate the Gs, Gi/o, and Gq/11 proteins and stimulate cAMP, inositol phosphates, and progesterone production, as well as ERK1/2 phosphorylation in the absence of agonist when expressed in the mouse Leydig MA-­10 tumor cells.683–685 Further, none of these mutants exhibited preferential or selective activation of any of these G proteins in basal or agonist-­ stimulated conditions, which suggests that the active receptor conformations provoked by these particular mutations did not lead to biased signaling of the mutant receptor, at least in the cell context of this particular tumor cell line.683 C. The Asp578His mutation has only been found in boys with gonadotropin-­independent precocious puberty-­bearing Leydig cell adenomas.648,686,687 This missense mutation causes Leydig cells to undergo a neoplastic transformation and, in contrast to all other LHCGR CAMs, which are germ-­line mutations, the Asp578His occurs somatically. It appears that the oncogenic effects of the Asp578His CAM are a particular feature of human Leydig cells and that these mitogenic effects are probably mediated by G protein-­ independent mechanisms.    Naturally occurring mutations of the FSHR are fewer in number but can be similarly classified. In fact, only ∼30 inactivating mutations in the hFSHR have been reported to date (Figs. 2.5 and 2.14).297,626,629,630,632 As it has been observed with the inactivating hLHCGR, there is in general a good correlation between residual activity exhibited by the mutant hFSHRs and the severity of the clinical phenotype expressed by the patients bearing the mutation(s).132,637,688 For example, the Ile423Thr mutant alters receptor activation, severely impacting ligand-­stimulated cAMP/ PKA signaling but much less so β-­arrestin-­associated ERK1/2 phosphorylation and led to premature ovarian failure. In the case of the Asp408Tyr mutant, which is a trafficking defective mutant exhibiting a poor agonist response to both endpoint measures, the phenotype in the affected women is severe congenital primary ovarian failure.626,689 The exception is the Phe591Ser mutation for which no clinical phenotype has been described; this particular mutation, which leads to severely impaired agonist binding and intracellular signaling, was found to be associated with the development of sex cord tumors.690 The most severe phenotype observed in homozygous females harboring inactivating mutations such as Ala189Val, Ala419Thr, or Pro519Ile is hypergonadotropic hypogonadism, arrest of follicular maturation beyond the primary stage, and complete lack of responsiveness to hFSH.623,637,691 Less severe phenotypes are observed in women who are homozygous or compound heterozygous for other mutations. These phenotypes may include secondary amenorrhea, gonadotropin resistance, and follicular development up to the antral stage.132,636 The phenotype in homozygous males is not clinically obvious as sperm quality is altered but fertility is preserved.635 The functional properties of many of these hFSHR mutants have been studied by expressing the mutants in heterologous cell lines. Ala419Thr in TM2 and Arg573Cys in the cytoplasmic face of TM6 impairs signal transduction while having little to no effect on hFSH binding or on the proper trafficking of the hFSHR to the PM.636,692 Mutant hFSHRs with functional defects of the Val221Gly (at the ectodomain)693 and Ala575Val (at the TMD6)694 have not been studied in detail, whereas in the case of the Pro348Arg hFSHR (located at the hinge region of the receptor), both ligand binding and agonist-stimulated sig naling were severely impaired.

TABLE 2.3  Functionally Characterized, Trafficking Defective hFHSR Mutations Region

Mutations

References

Extracellular domain or hinge region

Ile160Thr, Ala189Val, Asn191Ile, Asp224Val, and Met265Val Asp408Tyr

340, 636, 697, 698

Ala575Val, Leu597Ile Arg634His

699, 698 700

transmembrane domain 2 (TMD2) TMD6 Ctail

689

mutations interfere with the proper trafficking of the receptor from the ER to the PM is still unknown. As in the case of the hLHCGR, many loss-­of-­function mutations in the hFSHR are proteins with impaired transport of the receptor to the PM, thus inducing a complete lack of hFSH binding and therefore responsiveness to agonist.632,696 Among the mutant hFSHRs reported to date, at least ten are trafficking defective receptors which have been detected as intracellular retained molecules by imaging and/ or biochemical in vitro studies (Table 2.3). These mutations occur either at the ectodomain,340, 636, 697, 698 the TMD2,689 the EL2,691 the TMD6,699 and the Ctail.700 The naturally occurring mutation A189V caused a profound defect in targeting the receptor protein to the PM,701 confirming the importance of the 189Ala-­Phe-­Asn-­Gly-­Thr193 motif for intracellular trafficking of the hFSHR. This is also the locus of loss-­of-­function mutations in the hLHCGR646 (see above). Valine in the 189 position and isoleucine in position 191 may interfere with the structural integrity of the LRRs, which hosts the glycosylation site. Perturbation of this structure likely impairs proper receptor LRR formation, particularly its α-­helical portion. Although the putative loss of glycosylation may affect the folding and trafficking of the mutant receptor to the PM, it has yet to be determined whether this mutant form of the receptor is glycosylated or not at the Asn191 site. When the Ala189Val mutant is overexpressed in vitro, only a very small proportion of the mutated receptor is present at the PM but it is functional,624,701 albeit the FSH-­stimulated cAMP production by the PM-­expressed residual receptor is relatively higher than expected for its low membrane expression.623,701 Instead, most of the mutated receptor is sequestered inside the cell, explaining the inactivation mechanism.701 Interestingly, the reduced level of PM expression of the Ala189Val hFSHR confers preferential coupling to the β-­arrestin-­mediated signaling pathway, similar to that observed when the WT receptor is expressed at low PM densities, indicating that the selective signaling is due to the low level of PM expression rather than to the mutation itself.624 This observation might help to clarify why mutations of FSHβ are more deleterious to male fertility than the hFSHR Ala189Val mutation,151,635,702 which preserves a fraction of its receptor signaling repertoire. Other inactivating mutations in the hFSHR are also interesting. The Asn191Ile mutation was originally detected in the heterozygous state in a phenotypically normal women.697 Although the mutation affects the putative glycosylation site at position 191 of the receptor (see above), no studies on the effect of this mutation on the binding capacity and PM expression of the FSHR have been performed; in this regard, in vitro studies have been somehow contradictory regarding the impact of this mutation on agonist-­stimulated signaling.646,697 Nevertheless, they suggested that a fraction of the transfected receptor was, in fact, present at the PM. Given that glycosylation in only one position at the ectodomain is sufficient for folding and trafficking of the FSHR to the PM,456 these studies suggest that the limited PM expression of the mutant receptor may be mainly due to the alteration in the structural integrity of the Ala-­Phe-­Asn-­Gly-­Thr motif, rather than to the absence of glycosylation at this particular site. As mentioned above, the location of the mutation and the

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

nature of the amino acid substitution are both strong determinants of the functional features exhibited by mutant hFSHRs. For example, the Pro519Thr mutation in the center of the EL2 leads to complete failure to bind agonist and trigger intracellular signaling; women homozygous for this mutation express a severe phenotype including an absence of puberty, primary amenorrhea, and small ovaries.691 The effects of this particular mutation contrast with those provoked by other mutations in the serpentine region of the hFSHR (Ala419Thr, Arg573Cys, and Leu601Val), which usually results in partial receptor inactivation, with minimal effects on FSH binding.340,636,692 Therefore, it seems that the loss of a proline at position 519 provokes a severe conformational defect that leads to trapping of the receptor at the ER.691 Because the peptide backbone of proline is constrained in a ring structure, the occurrence of this amino acid is associated with a forced turn in the protein sequence, which is likely lost by the substitution with the more reactive threonine. Therefore, it is possible that the abrupt turn at the middle of the EL2 is a requisite not only for activity 338 but also for routing. In the case of the FSHR mutation at position Asp408Tyr (at the TMD2),689 molecular dynamics simulations have demonstrated that aspartate at this particular position is essential to maintain interhelical interactions between TMD2 and TMD7 and a conformational variability at the backbone level compatible with ER export to the PM. This is not the case with the Ile423Thr mutant, in which the replacement of isoleucine with threonine did not alter the interhelical connectivity between these two TMDs and thus did not disturb its normal trafficking to the PM.626,703 In contrast to the hLHCGR, only one “pure” hFSHR CAM (Asp567Gly, at the IL3-­ TMD6 junction, has thus far been detected in one male patient.704 This hFSHR CAM was identified in a man who exhibited normal spermatogenesis in a setting of undetectable circulating gonadotropins due to a previous hypophysectomy. Transgenic mice generated harboring the Asp567Gly mutation or FSHR WT on a gonadotropin deficient background have confirmed that this particular mutant behaves as a CAM in vivo and can stimulate a cAMP response in Sertoli cells and autonomous FSH-­ like actions in a ligand-­ deficient milieu.705–707 Other specific FSHR mutations are associated with spontaneous or pregnancy-­associated OHSS708–711 or impaired desensitization and internalization333 (Figs. 2.5, 2.14, and Table 2.2). In the OHSS cases, the mutant FSHRs exhibit a low level of constitutive activation as detected in vitro, but the OHSS phenotype is caused by the loss of ligand specificity of the mutated receptors, rendering them responsive to high levels of hCG or hTSH708–713 (see below). In the case of the Val514Ala mutation detected in a patient who developed OHSS during controlled ovarian hyperstimulation,699 the cell surface expression of the mutant FSHR in CHO cells was higher compared with the WT receptor, as was the production of cAMP and estradiol upon stimulation with low FSH doses in vitro and in vivo. Interestingly, the rate of FSH-­stimulated FSHR internalization was lower than that exhibited by the WT FSHR, which may have contributed to PM overexpression of the mutant receptor. Thus, it is possible that in this particular case, the OHSS developed as a consequence of the functional features of the Val514Ala FSHR rather than due to constitutive activity because neither the baseline levels of cAMP nor the response to hCG or TSH were different than those observed in the WT FSHR.699 Hence, the female phenotype of a genuine hFSHR CAM still remains unknown, and the pure male phenotype is based on only one patient. The Asn431Ile hFSHR CAM was serendipitously identified in a healthy man who exhibited normal spermatogenesis, suppressed serum FSH, and normal or elevated levels of biochemical markers for FSH action.333 It was particularly interesting that the mutation was located in the EL1, and the low level of constitutive activity (most probably due to reduced PM expression) was asso ciated with markedly altered agonist-

53

and internalization. Further studies with this particular mutant showed that the delayed desensitization and internalization of the mutant was probably due to failure to recruit endogenous β-­arrestin 1 properly.333 FSHR CAMs leading to spontaneous OHSS (Asp567Asn; Thr449Ala at TM3; and Ile545Thr at TMD5) are particularly interesting. These mutations lead to conformational changes in the receptor structure that, besides triggering modest constitutive activity, “relax” the binding specificity of the receptor, allowing the altered receptor to bind and become activated by high concentrations of hCG 708–711 or TSH.708–710 As discussed above, in the unliganded state the ectodomain of the hFSHR exerts a putative inhibitory influence on the TMD keeping the receptor in an inactive state. Upon agonist binding or when mutations are introduced at Ser273 (at the hinge region),318 the tethered extracellular loops of the TMDs are freed from this inhibitory influence allowing the TMDs to attain an active conformation.20 The pathogenesis of the OHSS associated with these CAMs has been explained by low-­affinity promiscuous interaction of hCG or TSH with the ectodomain of a mutant FSHR in a setting of a partially “unlocked” FSHR serpentine region, leading to excessive follicular recruitment.708 The ability of the hFSHR CAM Asp567Asn to become activated by hCG and TSH has also been confirmed in animal models.706 The rarity of naturally occurring hFSHR CAMs (four in total, Fig. 2.14) may have several explanations, such as:    A. The relative stability of the TMD of the hFSHR in the inactive state compared to the hLHCGR.714 In fact, basal cAMP production in hFSHR CAMs is quite modest and exposure to submaximal doses of hFSH usually results in a robust cAMP response.333,704,708–710 B.  The increased refractoriness of the hFSHR to mutation-­ induced constitutive activity compared to the hLHCGR.715 C. The lack of clearly defined phenotypes in both males and females that makes it difficult to identify naturally occurring hFSHR CAMs. D. Mutations in hFSHRs leading to constitutive activity could bias the activity toward either cAMP production, inositol triphosphate production or activation of β-­arrestin-­mediated pathways, such as ERK/MAPK signaling, complicating its identification unless all these endpoints are analyzed.

Polymorphisms of the Gonadotropin Receptor Genes At least 300 SNPs of the LHCGR and 2000 of the FSHR have been identified (http://www.ncbi.nlm.nih.gov/SNP/snp_ref. cgi?locusId=3973 and http://www.ncbi.nlm.nih.gov/SNP/snp_ ref.cgi?locusId=2492).166,716–719 Nevertheless, only a few of them are of clinical relevance. Only three SNPs of the LHCGR occur in exons. A six nucleotide in-­frame insertion/deletion between codons 18 and 19 of exon 1 (18insLQ) results in the expression of two hLHCGR variants that differ by the presence (insLQ) or absence of a Leu-­Gln pair near the NH2-­terminus of the mature receptor. In homozygous subjects, this variant has been associated with early-­onset breast cancer and short disease-­free survival in women.720–723 This receptor variant has a high prevalence in Caucasians716 and when expressed in vitro exhibited increased sensitivity to hCG and higher levels of PM expression than the non-­ LQ LHCGR variant.720 Two additional frequent SNPs in exon 10 of the hLHCGR gene code for either Asn or Ser in codons 291 and 312 in the hinge region of its ECD. Since the prevalence of the 312NS variant is high in male infertile patients, its presence has been associated with spermatogenic damage.724 The presence of Asn or Ser at either of these positions does not appear to change the PM expression of the hLHCGR,722 which is interesting given that codon 291 is a glycosylation site when Asn

2

54

PART I  The Fundamentals of Reproduction

Only five of the FSHR SNPs are exonic. They are all located in exon 10 but only four cause a coding change (Ala307Thr, Arg524Ser, Ala665Thr, and Ser680Asn). The most common and best studied are Ala307Thr and Ser680Asn (Fig. 2.5), which are expressed in a strong linkage disequilibrium. The most common allelic variants, Thr307/Asn680 and Ala307/Ser680, are almost equally distributed among Caucasians,717,718 but its occurrence is much lower in women of Hispanic origin.725 The Ala307Thr and Ser680Asn polymorphism appears to be associated with a lower response to exogenous hFSH when employed in controlled ovarian hyperstimulation protocols and therefore has a predictive value for determining the optimal dose of FSH to be used in controlled ovarian hyperstimulation protocols (women harboring the Ala307/Ser680 variant requiring higher doses of FSH.629,718) The Asn680Ser variant has also been associated with lower testicular volume in selected north European populations.726 In vitro studies have shown that the Ser680 variant exhibits altered signal transduction kinetics, increased β-­arrestin recruitment and agonist-­stimulated internalization, and decreased CREB phosphorylation, CREB-­dependent gene transcription, and nuclear PKA activation.727 A −29G>A polymorphism in the core promoter region of the FSHR has been associated with a reduction in the transcriptional activity of the receptor gene in women with the -­29A/A genotype, as well as with primary or secondary amenorrhea and poor response to exogenous FSH because of its effect on the expression of the FSHR.629 The minor allele frequency ranges from 50% to 70% in East Asia and Europe, respectively, and is extremely low in Hispanic women.725

Homologous Receptors As described above, GPHRs are characterized by the presence of a large NH2-­terminal ECD containing several LRRs that serves as a ligand-­binding site. The GPHR family is now included in the leucine-­rich repeat-­containing G-­protein-­coupled receptor (LGR) family, which has been expanded to include several LGRs grouped in Types A, B, and C. Type A LGRs include the GPHRs, Type B includes the LGR 4 to 6 (ligands are R-­spondins), and Type C includes LGR7 and 8 with the insulin-­ related peptides H2 relaxin and insulin-­like peptide 3 or INSL3 as cognate ligands.302 These latter LGRs are also known as the relaxin family peptide receptors (RXFP1 and RXFP2, respectively).728,729 LGRs are involved in diverse physiological processes, including reproduction, bone metabolism, cardio-­renal function, cell growth, and stem cell differentiation.730–735

Low Molecular Weight Gonadotropin Receptor Agonists and Antagonists Although the nature of the conformational changes in the 7TMD of the gonadotropin receptors following ligand binding is still unknown, the importance of the TMD in receptor activation is underscored by the constitutive activity triggered by mutations in this domain. Equally compelling are the analogous mutations in other well-­known GPCRs for which the mechanisms of activation have been characterized.350,352,353,355,357,736 This is despite the significant differences in ligand binding pocket location between receptors for small ligands and the gonadotropin receptors. In addition, the discovery of a number of low molecular weight (LMW) compounds with a molecular weight cutoff of 900 MW (which associates with good oral bioavailability) and that interact with the TMD of the gonadotropin receptors as positive (PAM) or negative (NAM) allosteric modulators, also have provided important clues on the mechanisms subserving gonadotropin receptor activation. The interest in these compounds is reflected by the fact that to date there are over 170 known LMW molecules targeting

GPHR. The binding of these LMW compounds is in contrast to the cognate ligands, which bind to the ECD of the receptor. It has long been a goal to replace the gonadotropin protein hormones with organic LMW small molecules that would stimulate gonadotropin receptors or modulate the activity of agonists, and that can be administered orally. Reduced molecular flexibility, as measured by the number of rotatable bonds, and low polar surface area or total hydrogen bond count (sum of donors and acceptors) are also found to be important predictors of good oral bioavailability, independent of molecular weight.737 Despite these predictors of good oral bioavailability, an increase in the activity of small molecules has been achieved with the preparation of bivalent, dimeric analogs; this approach has worked with other FSHR small molecules.738 This may reflect the unique nature of the FSHR, which appears less flexible than LHCGR and is less susceptible to activating mutations. A net increase in activity may be due to a decrease in off-­time having a greater effect than loss of bioavailability potential. Replacing gonadotropin hormones with small orally active molecules requires the identification of small-­molecule PAMs of LHCGR and FSHR with agonist activities. Moreover, it is critical that these not only activate the PKA and PKB pathways in a sensitive (in vitro nanomolar efficacy) and specific (minimal cross-­over with TSHR) manner.739–746 Initial small molecule agonists of FSHR did not appear to compete for FSH binding. Therefore, they are likely interacting with the TMDs to stabilize active conformers of FSHR. FSHR agonists will prove useful for induction of follicular growth and maturation, oocyte retrieval, or natural cycle conception. Recent efforts have resulted in promising potential FSHR agonists747–749 and LHCGR agonists that may be useful for treatment of undescended testis or induction of ovulation. For example, Org 43553750 exhibited a shorter circulatory half-­life than hCG, stimulated testosterone production in primary Leydig cells, and was able to induce ovulation in female mice and rats after a single-­dose oral administration without provoking OHSS in rodent models.751,752 Dimeric ligands based on the parent Org 43533 are also effective, allosteric LHCGR agonists.753 In this regard, allosteric LHCGR agonists may be administered for induction of thecal cell androgen production in cases where insufficient androgen is produced naturally, and as it is a substrate for estrogen production when FSH induces aromatase, a pure FSH agonist in the face of insufficient thecal androgen may not yield high-­quality oocytes.753 Another goal is to identify small molecules which inhibit gonadotropin action and which can be used as nonsteroidal contraceptives (block ovulation or block follicular development). Interestingly, substituent substitutions can convert small molecule PAM agonists of FSHR to NAMs.740,754 Taken at face value, those results suggest that orthosteric and allosteric binding pockets in the TMDs are spatially similar. For example, a thiazolidinone FSHR PAM agonist was modified so that it lost FSHR agonistic activity while gaining the ability to block FSH-­ induced cAMP and aromatase activity.740 One of the earliest studies of a selective nonpeptide antagonist of the FSHR was reported in 2002.755 This compound, identified by high-­throughput screening strategies, was a fairly weak inhibitor (low micromolar) probably because it was an allosteric modulator of binding, not a competitive inhibitor. This antagonist also inhibited FSH-­induced cAMP in CHO cells expressing FSH receptor, FSH-­induced progesterone in adrenal cells expressing FSH receptor, and FSH-­induced estrogen production in rat granulosa cells. Another FSHR antagonist was reported a few years later.754 In contrast to the former compound, this FSHR small molecule antagonist did not inhibit FSH binding to its receptor. However, it did block FSH-­induced cAMP production using human FSHR in CHO cells but with a 54-fold less potent inhibition of cAMP in a rat granulosa cell line. The compound was also effective in inhibiting follicle

CHAPTER 2  The Gonadotropin Hormones and Their Receptors

growth and ovulation in an ex vivo mouse culture. Interestingly this FSHR small molecule antagonist has recently been shown to have increased potency in vitro when made as a bivalent small molecule.738 Further structure activity-­guided medicinal chemistry performed with these antagonists may provide even greater potency.756 An FSHR NAM has been identified that blocks cAMP and progesterone production in rat granulosa cells without blocking estradiol production, demonstrating that the FSHR exhibits biased signaling, as described above.619 This molecule exhibited some weak LH receptor activity in addition to its FSH activity but had no TSH receptor activity. It was not completely effective in suppressing FSH-­induced follicle development and ovulation of oocytes likely due to the fact that it did not block estradiol production. Another FSHR NAM was subsequently discovered which blocks FSH-­induced cAMP as well as both FSH-­induced steroidogenic pathways (progesterone and estrogen).757 This molecule, ADX68692, is also active orally and therefore represents a potential orally active therapeutic for constitutively active FSH receptor and of course a nonsteroidal contraceptive, the latter a long-­sought goal of many women. It’s worth mentioning the dichotomy between FSHR and LHCGR. There are no small molecule antagonists of the LHCGR that have been developed and publicly disclosed. It may well be that the flexibility of the LHCGR still allows engagement of G-­proteins and subsequent activation of adenylate cyclase even when small molecules are bound. On the other hand, the FSH receptor that appears less flexible and therefore more stable to mutations, which might cause ligand-­independent activation, seems to have engendered a small number of small molecule inhibitor candidates, which are also active at the hLHCGR.627 As with other GPCRs,443 small membrane-­permeable molecules have also been employed experimentally as pharmacological chaperones to rescue the function of misfolded gonadotropin receptors. This is the case of Org 42599/Org 43553, which rescued the misfolded Ser616Tyr and Ala593Pro mutant LHCGRs leading to LCH,655 as well as Org41841, which rescued the function of the almost complete loss-­of-­function Ala189Val mutant FSHR758 in vitro. These allosteric modulators may be employed in the clinical arena to treat LCH, particularly the Type II variant, which is associated with mild clinical phenotypes and an improvement in the quality of sperm in men bearing mutant misfolded FSHRs. Recently, the effects of two small molecule FSHR allosteric activators on reprogramming trafficking and signaling at VEEs have been analyzed.759 A thiazolidinone derivative (T1)740,748,760 and a benzamide compound749 elicited distinct effects after stimulation of FSHR internalization; both cAMP increase and receptor recycling were higher with the former than with FSH alone. This occurred in an APPL1-­independent and dependent manner, respectively. The latter compound increased cAMP signaling and receptor recycling similar to and higher than FSH, respectively, and in an opposite APPL1-­regulated manner than in the case of the thiazolidinone compound.759 The distinctly different sensitivity of small allosteric ligands to the effects of APPL1 might eventually be exploited for clinical applications in assisted reproductive therapy. A major advance in glycoprotein hormone receptor structure and function was very recently made when the cryo-­electron

55

microscopy structures of CG occupied (ECD) active and inactive states (TM/Gsα) of the complete hormone receptor complex were determined.761 While certain binding modes of hCG were similar (i.e., CGα-­C-­terminal), some were different (CGβ-­C-­terminal compared) to FSH/FSHR interaction with the ECD alone.314 The differences between FSHβ and CGβ C-­terminal binding to their respective cognate ECDs were ascribed to underpinning the specificity of binding. Notably for the first time, the full-­length LHR structure allowed for the determination of a major shift of the ECD by a 45° angle away from the TM domains, inferring movement away from the plasma membrane in the active state and rigidity of the complex, which would inhibit oligomerization. Additionally, the determination of a small molecule LHR agonist bound in the activated complex will allow for structure-­based drug design of new orally active molecules that activate or inhibit the receptor function. TOP REFERENCES

1. Hearn MTW, Gomme PT. Molecular architecture and biorecognition processes of the cystine knot protein superfamily: part I. The glycoprotein hormones. J Mol Recognit. 2000;13(5):223–278. 2. Lapthorn AJ, Harris DC, Littlejohn A. Crystal-­Structure of human Chorionic-­Gonadotropin. Nature. 1994;369:455–461. 3. Wu H, Lustbader JW, Liu Y. Structure of human Chorionic-­ Gonadotropin at 2.6-­angstrom resolution from mad analysis of the selenomethionyl protein. Structure. 1994;2:545–558. 4. Puett D, Wu C, Narayan P. The tie that binds: design of biologically active single-­ chain human chorionic gonadotropins and a gonadotropin-­ receptor complex using protein engineering. Biol Reprod. 1998;58:1337–1342. 5. Fox KM, Dias JA, Van Roey P. Three-­dimensional structure of human follicle-­stimulating hormone. Mol Endocrinol. 2001;15(3):378–389. 6. Bousfield GR, Harvey DJ. Follicle-­stimulating hormone glycobiology. Endocrinology. 2019;160:1515–1535. 7. Stamatiades GA, Kaiser UB. Gonadotropin regulation by pulsatile GnRH: signaling and gene expression. Mol Cell Endocrinol. 2018;463:131–141. 8. Ascoli M, Fanelli F, Segaloff DL. The lutropin/choriogonadotropin receptor. A 2002 perspective. Endocr Rev. 2002;23:141–174. 9. Richards JS. From follicular development and ovulation to ovarian cancers: an unexpected journey. Vitam Horm. 2018;47(107):453–472. 10. Ulloa-­Aguirre A, Reiter E, Crépieux P. FSH receptor signaling: complexity of interactions and signal diversity. Endocrinology. 2018;159(8):3020–3035. 11. Ulloa-­Aguirre A, Reiter E, Bousfield G, Dias JA, Huhtaniemi I. Constitutive activity in gonadotropin receptors. Adv Pharmacol. 2014;70:37–80. 12. Brüser A, Schulz A, Rothemund S, et al. The activation mechanism of glycoprotein hormone receptors with implications in the cause and therapy of endocrine diseases. J Biol Chem. 2016;291(2):508–520. 13. Jiang X, Dias JA, He X. Structural biology of glycoprotein hormones and their receptors: insights to signaling. Mol Cell Endocrinol. 2014;382(1):424–451. 14. Ulloa-­Aguirre A, Zariñán T, Dias JA, Conn PM. Mutations in G protein-­coupled receptors that impact receptor trafficking and reproductive function. Mol Cell Endocrinol. 2014;382(1):411–423. 15. Landomiel F, De Pascali F, Raynaud P, et al. Biased signaling and allosteric modulation at the FSHR. Front Endocrinol (Lausanne). 2019;10:148.

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REFERENCES

1. Hearn MTW, Gomme PT. Molecular architecture and biorecognition processes of the cystine knot protein superfamily: part I. The glycoprotein hormones. J Mol Recognit. 2000;13(5):223–278. 2. Pierce JG, Parsons TF. Glycoprotein hormones: structure and function. Annu Rev Biochem. 1981;50:465–495. 3. Huhtaniemi IT, Clayton RN, Catt KJ. Gonadotropin binding and Leydig cell activation in the rat testis in vivo. Endocrinology. 1982;111(3):982–987. 4. Dias JA, Cohen BD, Lindau-­Shepard B, Nechamen CA, Peterson AJ, Schmidt A. Molecular, structural, and cellular biology of follitropin and follitropin receptor. Vitam Horm. 2002;64:249–322. 5. Richards JS, Ascoli M. Endocrine, paracrine, and autocrine signaling pathways that regulate ovulation. Trends Endocrinol Metab. 2018;29(5):313–325. 6. Richards JS. Hormonal control of gene expression in the ovary. Endocr Rev. 1994;15(6):725–751. 7. Richards JS, Pangas SA. The ovary: basic biology and clinical implications. J Clin Invest. 2010;120(4):963–972. 8. Banerjee P, Fazleabas AT. Extragonadal actions of chorionic gonadotropin. Rev Endocr Metab Disord. 2011;12(4):323–332. 9. Fournier T, Guibourdenche J, Evain-­Brion D. Review: hCGs: different sources of production, different glycoforms and functions. Placenta. 2015;36(suppl 1):S60–S65. 10. Huhtaniemi IT, Korenbrot CC, Jaffe RB. HCG binding and stimulation of testosterone biosynthesis in the human fetal testis. J Clin Endocrinol Metab. 1977;44(5):963–967. 11. Fiddes JC, Goodman HM. The CDNA for the beta-­subunit of human chorionic-­gonadotropin suggests evolution of a gene by readthrough into the 3’-­untranslated region. Nature. 1980;286(5774):684–687. 12. Nakav S, Jablonka-­Shariff A, Kaner S, Chadna-­Mohanty P, Grotjan HE, Ben-­Menahem D. The LH beta gene of several mammals embeds a carboxyl-­ terminal peptide-­ like sequence revealing a critical role for mucin oligosaccharides in the evolution of lutropin to chorionic gonadotropin in the animal phyla. J Biol Chem. 2005;280(17):16676–16684. 13. Talmadge K, Boorstein WR, Vamvakopoulos NC, Gething MJ, Fiddes JC. Only 3 of the 7 human chorionic-­gonadotropin beta-­ subunit genes can be expressed in the placenta. Nucleic Acids Res. 1984;12(22):8415–8436. 14. Talmadge K, Vamvakopoulos NC, Fiddes JC. Evolution of the genes for the beta-­subunits of human chorionic-­gonadotropin and luteinizing-­hormone. Nature. 1984;307(5946):37–40. 15. Lapthorn AJ, Harris DC, Littlejohn A. Crystal-­structure of human chorionic-­gonadotropin. Nature. 1994;369:455–461. 16. Wu H, Lustbader JW, Liu Y. Structure of human chorionic-­ gonadotropin at 2.6-­angstrom resolution from mad analysis of the Selenomethionyl protein. Structure. 1994;2:545–558. 17. Tegoni M, Spinelli S, Verhoeyen M, Davis P, Cambillau C. Crystal structure of a ternary complex between human chorionic gonadotropin (hCG) and two Fv fragments specific for the alpha and beta-­ subunits. J Mol Biol. 1999;289(5):1375–1385. 18. Fox KM, Dias JA, Van Roey P. Three-­ dimensional structure of human follicle-­ stimulating hormone. Mol Endocrinol. 2001;15(3):378–389. 19. Fan QR, Hendrickson WA. Structure of human follicle-­ stimulating hormone in complex with its receptor. Nature. 2005;433(7023):269–277. 20. Jiang X, Liu H, Chen X, et al. Structure of follicle-­stimulating hormone in complex with the entire ectodomain of its receptor. Proc Natl Acad Sci U S A. 2012;109(31):12491–12496. 21. Larson SB, McPherson A. The crystal structure of the beta subunit of luteinizing hormone and a model for the intact hormone. Curr Res Struct Biol. 2019;1:1–5. 22. DeBeer T, VanZuylen C, Leeflang BR, et al. NMR studies of the free alpha subunit of human chorionic gonadotropin—structural influences of N-­glycosylation and the beta subunit on the conformation of the alpha subunit. Eur J Biochem. 1996;241(1): 229–242. 23. Erbel PJ, Karimi-­Nejad Y, De Beer T, Boelens R, Kamerling JP, Vliegenthart JF. Solution structure of the alpha-­subunit of human chorionic gonadotropin. Eur J Biochem. 1999;260(2):490–498. 24. Erbel PJA, Haseley SR, Kamerling JP, Vliegenthart JFG. Studies on the relevance of the glycan at Asn-52 of the alpha-subunit of human chorionic gonadotropin in the alpha2002;364:485–495.

25. Heikoop JC, Van Den Boogaart P, De Leeuw R, Rose UM, Mulders JWM, Grootenhuis PDJ. Partially deglycosylated human choriogonadotropin, stabilized by intersubunit disulfide bonds, shows full bioactivity. Eur J Biochem. 1998;253(1):354–356. 26. Cole LA, Khanlian SA. Hyperglycosylated hCG: a variant with separate biological functions to regular hCG. Mol Cell Endocrinol. 2007;260:228–236. 27. Fares F. The role of O-­linked and N-­linked oligosaccharides on the structure-­function of glycoprotein hormones: development of agonists and antagonists. Biochim Biophys Acta. 2006;1760(4):560–567. 28. Ulloa-­Aguirre A, Maldonado A, Damian-­Matsumura P, Timossi C. Endocrine regulation of gonadotropin glycosylation. Arch Med Res. 2001;32(6):520–532. 29. Jiang C, Hou X, Wang C, et al. Hypoglycosylated hFSH has greater bioactivity than fully glycosylated recombinant hFSH in human granulosa cells. J Clin Endocrinol Metab. 2015;100(6):E852–E860. 30. Bousfield GR, May JV, Davis JS, Dias JA, Kumar TR. In vivo and in vitro impact of carbohydrate variation on human follicle-­ stimulating hormone function. Front Endocrinol. 2018;9:216. 31. Liang A, Plewes MR, Hua G, et al. Bioactivity of recombinant hFSH glycosylation variants in primary cultures of porcine granulosa cells. Mol Cell Endocrinol. 2020;514:110911. 32. Wang H, May J, Butnev V, et al. Evaluation of in vivo bioactivities of recombinant hypo-­(FSH21/18) and fully-­(FSH24) glycosylated human FSH glycoforms in Fshb null mice. Mol Cell Endocrinol. 2016;437:224–236. 33. Zarinan T, Butnev VY, Gutierrez-­Sagal R, et al. In vitro impact of FSH glycosylation variants on FSH receptor-­stimulated signal transduction and functional selectivity. J Endocr Soc. 2020;4(5). bvaa019. 34. Simon LE, Liu Z, Bousfield GR, Kumar TR, Duncan FE. Recombinant FSH glycoforms are bioactive in mouse preantral ovarian follicles. Reproduction. 2019;158(6):517–527. 35. Bousfield GR, Butnev VY, Gotschall RR, Baker VL, Moore WT. Structural features of mammalian gonadotropins. Mol Cell Endocrinol. 1996;125(1–2):3–19. 36. Dalpathado DS, Irungu J, Go EP, et al. Comparative glycomics of the glycoprotein follicle stimulating hormone: glycopeptide analysis of isolates from two mammalian species. Biochemistry. 2006;45(28):8665–8673. 37. Weisshaar G, Hiyama J, Renwick AGC, Nimtz M. NMR investigations of the n-­linked oligosaccharides at individual glycosylation sites of human lutropin. Eur J Biochem. 1991;195(1):257–268. 38. Baenziger, JU. Protein-­specific glycosyltransferases—how and why they do it. FASEB J. 1994;8(13):1019–1025. 39. Mi Y, Fiete D, Baenziger, JU. Ablation of GalNAc-­4-­sulfotransferase-­1 enhances reproduction by altering the carbohydrate structures of luteinizing hormone in mice. J Clin Invest. 2008;118(5):1815–1824. 40. Green ED, Baenziger, JU. Asparagine-­linked oligosaccharides on lutropin, follitropin, and thyrotropin.1. Structural elucidation of the sulfated and sialylated oligosaccharides on bovine, ovine, and human pituitary glycoprotein hormones. J Biol Chem. 1988;263(1):25–35. 41. Green ED, Baenziger JU. Asparagine-­linked oligosaccharides on lutropin, follitropin, and thyrotropin. 2. Distributions of sulfated and sialylated oligosaccharides on bovine, ovine, and human pituitary glycoprotein hormones. J Biol Chem. 1988;263(1):36–44. 42. Bousfield GR, Butnev VY, Rueda-­ Santos MA, Brown A, Hall AS, Harvey DJ. Macro-­and micro-­heterogeneity in pituitary and urinary follicle-­ stimulating hormone glycosylation. J Glycomics Lipidomics. 2014;4:1000125. 43. Bousfield GR, Butnev VY, Walton WJ, et al. All-­ or-­ none N-­ glycosylation in primate follicle-­ stimulating hormone beta-­ subunits. Mol Cell Endocrinol. 2007;260–262:40–48. 44. Bousfield GR, Dias JA. Synthesis and secretion of gonadotropins including structure-­function correlates. Rev Endocr Metab Disord. 2011;12(4):289–302. 45. Liu CL, Bowers LD. Mass spectrometric characterization of the beta-­subunit of human chorionic gonadotropin. J Mass Spectrom. 1997;32(1):33–42. 46. Toll H, Berger P, Hofmann A, et al. Glycosylation patterns of human chorionic gonadotropin revealed by liquid chromatography-­mass spectrometry and bioinformatics. Electrophoresis. 2006;27(13):2734–2746. 47. Valmu L, Alfthan H, Hotakainen K, Birken S, Stenman UH. Site-­ specific glycan analysis of human chorionic gonadotropin beta-­ subunit from malignancies and pregnancy by liquid electrospray mass spectrometry. Glycobiology.

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PART I  The Fundamentals of Reproduction

48. Darling RJ, Ruddon RW, Perini F, Bedows E. Cystine knot mutations affect the folding of the glycoprotein hormone alpha-­subunit -­differential secretion and assembly of partially folded intermediates. J Biol Chem. 2000;275(20):15413–15421. 49. Darling RJ, Wilken JA, Miller-­ Lindbolm AK, et al. Functional contributions of noncysteine residues within the cystine knots of human chorionic gonadotropin subunits. J Biol Chem. 2001;276(14):10692–10699. 50. Darling RJ, Wilken JA, Ruddon RW, Bedows E. Intracellular folding pathway of the cystine knot-­containing glycoprotein hormone alpha-­subunit. Biochemistry. 2001;40(2):577–585. 51. Wilken JA, Bedows E. Disuflide bond rearrangement during formation of the chorionic gonadotropin beta-­subunit cystine knot in vivo. Biochemistry. 2004;43(17):5109–5118. 52. Wilken JA, Bedows E. A novel four-­amino acid determinant defines conformational freedom within chorionic gonadotropin beta-­ subunits. Biochemistry. 2007;46(14):4417–4424. 53. Xing YN, Myers RV, Cao DH, et al. Glycoprotein hormone assembly in the endoplasmic reticulum—I. The glycosylated end of human alpha-­subunit loop 2 is threaded through a beta-­subunit hole. J Biol Chem. 2004;279(34):35426–35436. 54. Xing YN, Myers RV, Cao DH, et al. Glycoprotein hormone assembly in the endoplasmic reticulum—II. Multiple roles of a redox sensitive beta-­ subunit disulfide switch. J Biol Chem. 2004;279(34):35437–35448. 55. Xing YN, Myers RV, Cao DH, et al. Glycoprotein hormone assembly in the endoplasmic reticulum—III. The seatbelt and its latch site determine the assembly pathway. J Biol Chem. 2004;279(34):35449–35457. 56. Xing YN, Myers RV, Cao DH, et al. Glycoprotein hormone assembly in the endoplasmic reticulum—IV. Probable mechanism of subunit docking and completion of assembly. J Biol Chem. 2004;279(34):35458–35468. 57. Xing YN, Williams C, Campbell RK, et al. Threading of a glycosylated protein loop through a protein hole: implications for combination of human chorionic gonadotropin subunits. Protein Sci. 2001;10(2):226–235. 58. Krause JM, Berger P, Roig J, Singh V, Merz WE. Rapid maturation of glycoprotein hormone free alpha-­ subunit (GPHalpha) and GPHalpha alpha homodimers. Mol Endocrinol. 2007;21(10):2551–2564. 59. Merz WE, Krause JM, Roig J, Singh V, Berger P. Non-­assembled human chorionic gonadotropin subunits and αα-­homodimers use fast-­track processing in the secretory pathway in contrast to αβ-­ heterodimers. Endocrinology. 2007;148(12):5831–5841. 60. Roig J, Krause JM, Berger P, Merz WE. Time-­dependent folding of immunological epitopes of the human chorionic gonadotropin beta-­subunit. Mol Cell Endocrinol. 2007;260:12–22. 61. Singh V, Merz WE. Disulfide bond formation is not required for human chorionic gonadotropin subunit association—studies with dithiothreitol in JEG-­3 cells. J Biol Chem. 2000;275(16):11765–11770. 62. Bernard MP, Lin W, Kholodovych V, Moyle WR. Human lutropin (hLH) and choriogonadotropin (CG) are assembled by different pathways: a model of hLH assembly. J Biol Chem. 2014;289(20):14360–14369. 63. Butler SA, Laidler P, Porter JR, et al. The beta-­subunit of human chorionic gonadotrophin exists as a homodimer. J Mol Endocrinol. 1999;22(2):185–192. 64. Peng KC, Puett D, Brewer JM. Homodimer formation by the individual subunits of bovine lutropin as determined by sedimentation equilibrium. J Mol Endocrinol. 1997;18(3):259–265. 65. Berger P, Gruschwitz M, Spoettl G, et al. Human chorionic gonadotropin (hCG) in the male reproductive tract. Mol Cell Endocrinol. 2007;260:190–196. 66. Nemansky M, Moy E, Lyons CD, Yu I, Blithe DL. Human endometrial stromal cells generate uncombined alpha-­subunit from human chorionic gonadotropin, which can synergize with progesterone to induce decidualization. J Clin Endocrinol Metab. 1998;83(2):575–581. 67. Szkudlinski MW, Teh NG, Grossmann M, Tropea JE, Weintraub BD. Engineering human glycoprotein hormone superactive analogues. Nat Biotechnol. 1996;14(10):1257–1263. 68. Shao K, Purohit S, Bahl OP. Effect of modification of all loop regions in the alpha-­and beta-­ subunits of human choriogonadotropin on its signal transduction activity. 1996;122(2):173–182.

69. Thotakura NR, Blithe DL. Glycoprotein hormones—glycobiology of gonadotropins, thyrotropin and free alpha-­subunit. Glycobiology. 1995;5(1):3–10. 70. Krystek SR, Dias JA, Andersen TT. Identification of a subunit contact site of the alpha-­ subunit of follitropin. Pept Res. 1992;5(3):165–168. 71. Liu C, Roth KE, Shepard BAL, Shaffer JB, Dias JA. Site-­Directed alanine mutagenesis of Phe(33), Arg(35), and Arg(42)-­ Ser(43)-­ Lys(44) in the human gonadotropin alpha-­subunit. J Biol Chem. 1993;268(29):21613–21617. 72. Xia HY, Chen F, Puett D. A region in the human glycoprotein hormone alpha-­subunit important in holoprotein formation and receptor-­binding. Endocrinology. 1994;134(4):1768–1770. 73. Chen F, Wang Y, Puett D. The carboxy-­terminal region of the glycoprotein hormone alpha-­subunit -­contributions to receptor-­ binding and signaling in human chorionic-­ gonadotropin. Mol Endocrinol. 1992;6(6):914–919. 74. Yoo JK, Zeng HW, Ji IH, Murdoch WJ, Ji TH. COOH-­terminal amino acids of the alpha subunit play common and different roles in human choriogonadotropin and follitropin. J Biol Chem. 1993;268(18):13034–13042. 75. Chen F, Wang Y, Puett D. Role of the invariant aspartic acid-­ 99 of human choriogonadotropin-­ beta in receptor-­ binding and biological-­activity. J Biol Chem. 1991;266(29):19357–19361. 76. Hu SB, Johnson L, Roche PC, Keutmann HT. A functional determinant in human luteinizing hormone and chorionic gonadotropin: differential effect of mutations about beta-­GLN-­S4. Endocrinology. 1997;138(4):1627–1633. 77. Huang JN, Chen F, Puett D. Amino carboxyl-­ terminal deletion mutants of human choriogonadotropin-­ beta. J Biol Chem. 1993;268(13):9311–9315. 78. Huang JN, Puett D. On the role of the invariant glutamine at position-­54 in the human choriogonadotropin beta-­subunit. Mol Cell Biochem. 1994;136(2):183–186. 79. Matzuk MM, Hsueh AJW, Lapolt P, Tsafriri A, Keene JL, Boime I. The biological role of the carboxyl-­terminal extension of human chorionic-­ gonadotropin beta-­ subunit. Endocrinology. 1990;126(1):376–383. 80. Morbeck DE, Roche PC, Keutmann HT, McCormick DJ. A receptor-­ binding site identified in the region 81–95 of the beta-­ subunit of human luteinizing-­ hormone (LH) and chorionic-­ gonadotropin (HCG). Mol Cell Endocrinol. 1993;97(1–2):173–181. 81. Xia HY, Fernandez LM, Puett D. Replacement of the invariant tyrosine in the cagy region of the human chorionic-­gonadotropin beta subunit. Mol Cell Endocrinol. 1993;92(1):R1–R5. 82. Xia HY, Puett D. Identification of conserved amino-­acid-­residues in the beta-­subunit of human choriogonadotropin important in holoprotein formation. J Biol Chem. 1994;269(27):17944–17953. 83. Eldeiry S, Kaetzel D, Kennedy G, Nilson J, Puett D. Site-­directed mutagenesis of the human chorionic-­gonadotropin beta-­subunit— bioactivity of a heterologous hormone, bovine alpha-­human DES-­ (122–145)-­beta. J Mol Endocrinol. 1989;3(10):1523–1528. 84. Campbell RK, Bergert ER, Wang YH, Morris JC, Moyle WR. Chimeric proteins can exceed the sum of their parts: implications for evolution and protein design. Nat Biotechnol. 1997;15(5):439–443. 85. Cosowsky L, Lin W, Han Y, Bernard MP, Campbell RK, Moyle WR. Influence of subunit interactions on lutropin specificity— implications for studies of glycoprotein hormone function. J Biol Chem. 1997;272(6):3309–3314. 86. Dias JA, Zhang YQ, Liu XX. Receptor-­binding and functional-­ properties of chimeric human follitropin prepared by an exchange between a small hydrophilic intercysteine loop of human follitropin and human lutropin. J Biol Chem. 1994;269(41):25289–25294. 87. Grossmann M, Szkudlinski MW, Wong R, Dias JA, Ji TH, Weintraub BD. Substitution of the seat-­ belt region of the thyroid-­ stimulating hormone (TSH) beta-­ subunit with the corresponding regions of choriogonadotropin or follitropin confers luteotropic but not follitropic activity to chimeric TSH. J Biol Chem. 1997;272(24):15532–15540. 88. Han Y, Bernard MP, Moyle WR. hCG beta residues 94–96 alter LH activity without appearing to make key receptor contacts. Mol Cell Endocrinol. 1996;124(1–2):151–161. . The lutropin beta-­ terminus facilitates subunit combination by offsetting

CHAPTER 2  The Gonadotropin Hormones and Their Receptors the inhibitory effects of residues needed for LH activity. Mol Cell Endocrinol. 1995;112(1):21–25. 90. Wang YH, Bernard MP, Moyle WR. Bifunctional hCG analogs adopt different conformations in LH and FSH receptor complexes. Mol Cell Endocrinol. 2000;170(1–2):67–77. 91. Ben-­Menahem D. Single chain variants of the glycoprotein hormones and their receptors as tools to study receptor activation and for analogue design. J Neuroendocrinol. 2004;16(2):171–177. 92. Narayan P, Gray J, Puett D. A biologically active single chain human chorionic gonadotropin analog with altered receptor binding properties. Endocrinology. 2000;141(1):67–71. 93. Narayan P, Wu CB, Puett D. Functional expression of yoked human chorionic-­gonadotropin in baculovirus-­infected insect cells. J Mol Endocrinol. 1995;9(12):1720–1726. 94. Setlur SR, Dighe RR. Single chain human chorionic gonadotropin, hCG alpha beta: effects of mutations in the alpha subunit on structure and bioactivity. Glycoconj J. 2007;24(1):97–106. 95. Sugahara T, Pixley MR, Minami S, et al. Biosynthesis of a biologically-­active single peptide-­chain containing the human common alpha-­subunits and chorionic-­gonadotropin beta-­subunits in tandem. Proc Natl Acad Sci U S A. 1995;92(6):2041–2045. 96. BenMenahem D, Kudo M, Pixley MB, et al. The biologic action of single-­ chain choriogonadotropin is not dependent on the individual disulfide bonds of the beta subunit. J Biol Chem. 1997;272(11):6827–6830. 97. Sato A, Perlas E, BenMenahem D, et al. Cystine knot of the gonadotropin alpha subunit is critical for intracellular behavior but not for in vitro biological activity. J Biol Chem. 1997;272(29):18098–18103. 98. GarciaCampayo V, Sato A, Hirsch B, et al. Design of stable biologically active recombinant lutropin analogs. Nat Biotechnol. 1997;15(7):663–667. 99. Sugahara T, Grootenhuis PDJ, Sato A, et al. Expression of biologically active fusion genes encoding the common alpha subunit and either the CG beta or FSH beta subunits: role of a linker sequence. Mol Cell Endocrinol. 1996;125(1–2):71–77. 100. Bernard MP, Lin W, Myers R, Cao D, Xing Y, Moyle WR. Crosslinked bifunctional gonadotropin analogs with reduced efficacy. Mol Cell Endocrinol. 2005;233(1–2):25–31. 101. Einstein M, Lin W, Macdonald GJ, Moyle WR. Partial restoration of lutropin activity by an intersubunit disulfide bond: implications for structure/function studies. Exp Biol Med. 2001;226(6):581–590. 102. Heikoop JC, Huisman-­de Winkel B, Grootenhuis PDJ. Towards minimized gonadotropins with full bioactivity. Eur J Biochem. 1999;261(1):81–83. 103. Heikoop JC, vanBeuningendeVaan M, vandenBoogaart P, Grootenhuis PDJ. Evaluation of subunit truncation and the nature of the spacer for single chain human gonadotropins. Eur J Biochem. 1997;245(3):656–662. 104. Heikoop JC, vandenBoogaart P, Mulders JWM, Grootenhuis PDJ. Structure-­based design and protein engineering of intersubunit disulfide bonds in gonadotropins. Nat Biotechnol. 1997;15(7):658–662. 105. Arnold CJ, Liu C, Lindau-­Shepard B, Losavio ML, Patrascu MT, Dias JA. The human follitropin alpha-­subunit C terminus collaborates with a beta-­subunit cystine noose and an alpha-­subunit loop to assemble a receptor-­binding domain competent for signal transduction. Biochemistry. 1998;37(7):1762–1768. 106. Roy S, Setlur S, Gadkari RA, Krishnamurthy HN, Dighe RR. Translational fusion of two beta-­ subunits of human chorionic gonadotropin results in production of a novel antagonist of the hormone. Endocrinology. 2007;148(8):3977–3986. 107. Vardhana PA, Julius MA, Pollak SV, Lustbader EG, Trousdale RK, Lustbader JW. A unique human chorionic gonadotropin antagonist suppresses ovarian hyperstimulation syndrome in rats. Endocrinology. 2009;150(8):3807–3814. 108. Garcia-­Campayo V, Boime I. Independent activities of FSH and LH structurally confined in a single polypeptide: selective modification of the relative potencies of the hormones. Endocrinology. 2001;142(12):5203–5211. 109. Garcia-­Campayo V, Jablonka-­Shariff A, Boime I. A single-­chain bifunctional gonadotropin analog is secreted from Chinese hamster ovary cells as two distinct bioactive species. J Biol Chem. 2004;279(43):44286–44293. 110. Jablonka-­Shariff A, Comstock A, Colgin M, Adams T, Boime I. Construction and expression of an ovine single chain triple-­domain chimeric gonadotropin: tandem linkage of the genes encoding the alpha-­and FSH beta and LH beta subunits. Biol Reprod Reprod. 2007;77 (Issue Suppl_1):131–132.

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111. Kanda M, Jablonka-­Shariff A, Sato A, et al. Genetic fusion of an alpha-­subunit gene to the follicle-­stimulating hormone and chorionic gonadotropin-­beta subunit genes: production of a bifunctional protein. J Mol Endocrinol. 1999;13(11):1873–1881. 112. Jablonka-­Shariff A, Kumar TR, Eklund J, Comstock A, Boime I. Single-­ chain, triple-­ domain gonadotropin analogs with disulfide bond mutations in the alpha-­subunit elicit dual follitropin and lutropin activities in vivo. Mol Endocrinol. 2006;20(6):1437–1446. 113. Garcia-­Campayo V, Boime I, Ma XP, Daphna-­Iken D, Kumar TR. A single-­ chain tetradomain glycoprotein hormone analog elicits multiple hormone activities in vivo. Biol Reprod. 2005;72(2):301–308. 114. Garcia-­Campayo V, Kumar TR, Boime I. Thyrotropin, follitropin, and chorionic gonadotropin expressed as a single multifunctional unit reveal remarkable permissiveness in receptor-­ligand interactions. Endocrinology. 2002;143(10):3773–3778. 115. Meehan TP, Harmon BG, Overcast ME, et al. Gonadal defects and hormonal alterations in transgenic mice expressing a single chain human chorionic gonadotropin-­lutropin receptor complex. J Mol Endocrinol. 2005;34(2):489–503. 116. Meehan TP, Narayan P. Constitutively active luteinizing hormone receptors: consequences of in vivo expression. Mol Cell Endocrinol. 2007;260–262:294–300. 117. Narayan P, Wu CB, Puett D. Genetic engineering of single-­chain gonadotropins and hormone-­ receptor fusion proteins. Methods. 2000;21(1):59–66. 118. Puett D, Wu C, Narayan P. The tie that binds: design of biologically active single-­ chain human chorionic gonadotropins and a gonadotropin-­ receptor complex using protein engineering. Biol Reprod. 1998;58:1337–1342. 119. Wu CB, Narayan P, Puett D. Protein engineering of a novel constitutively active hormone-­ receptor complex. J Biol Chem. 1996;271(49):31638–31642. 120. Narayan P, Gray J, Puett D. Yoked complexes of human choriogonadotropin and the lutropin receptor: evidence that monomeric individual subunits are inactive. J Mol Biol. 2002;16(12):2733–2745. 121. Henke A, Gromoll J. New insights into the evolution of chorionic gonadotrophin. Mol Cell Endocrinol. 2008;291(1–2):11–19. 122. Nagirnaja L, Rull K, Uuskula L, Hallast P, Grigorova M, Laan M. Genomics and genetics of gonadotropin beta-­subunit genes: unique FSHB and duplicated LHB/CGB loci. Mol Cell Endocrinol. 2010;329(1–2):4–16. 123. Sherman GB, Wolfe MW, Farmerie TA, et al. A single gene encodes the beta-­subunits of equine luteinizing hormone and chorionic gonadotropin. Mol Endocrinol. 1992;6(6):951–959. 124. Policastro PF, Danielsmcqueen S, Carle G, Boime I. A map of the hCG-­beta-­LH-­beta gene-­cluster. J Biol Chem. 1986;261(13):5907–5916. 125. Gharib SD, Wierman ME, Shupnik MA, Chin WW. Molecular-­ biology of the pituitary gonadotropins. Endocr Rev. 1990;11(1):177–199. 126. Rull K, Laan M. Expression of beta-­subunit of HCG genes during normal and failed pregnancy. Hum Reprod. 2005;20(12):3360–3368. 127. Dirnhofer S, Hermann M, Hittmair A, Hoermann R, Kapelari K, Berger P. Expression of the human chorionic gonadotropin-­beta gene cluster in human pituitaries and alternate use of exon 1. J Clin Endocrinol Metab. 1996;81(12):4212–4217. 128. Berger P, Kranewitter W, Madersbacher S, Gerth R, Geley S, Dirnhofer S. Eutopic production of human chorionic-­gonadotropin-­ beta (hCG-­beta) and luteinizing-­hormone-­beta (hLH-­beta) in the human testis. FEBS Lett. 1994;343(3):229–233. 129. Giovangrandi Y, Parfait B, Asheuer M, et al. Analysis of the human CGB/LHB gene cluster in breast tumors by real-­time quantitative RT-­PCR assays. Cancer Lett. 2001;168(1):93–100. 130. Strauss BL, Pittman R, Pixley MR, Nilson J, Boime I. Expression of the beta-­subunit of chorionic-­gonadotropin in transgenic mice. J Biol Chem. 1994;269(7):4968–4973. 131. Nishimura R, Shin J, Ji I, et al. A single amino acid substitution in an ectopic alpha subunit of a human carcinoma choriogonadotropin. J Biol Chem. 1986;261(23):10475–10477. 132. Huhtaniemi IT, Themmen AP. Mutations in human gonadotropin and gonadotropin-­receptor genes. Endocrine. 2005;26(3):207–217. 133. Themmen APN. An update of the pathophysiology of human gonadotrophin subunit and receptor gene mutations and polymorphisms. Reproduction. 2005;130(3):263–274. 134. Weiss J, Axelrod L, Whitcomb RW, Harris PE, Crowley WF, . Hypogonadism caused by a single amino-­acid substitution in the beta subunit of luteinizing-hormone. N Engl J Med. 1992;326(3):179–183.

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PART I  The Fundamentals of Reproduction

135. Valdes-­Socin H, Salvi R, Daly AF, et al. Brief report: hypogonadism in a patient with a mutation in the luteinizing hormone beta-­subunit gene. N Engl J Med. 2004;351(25):2619–2625. 136. Lofrano-­Porto A, Barra GB, Giacomini LA, et al. Luteinizing hormone beta mutation and hypogonadism in men and women. N Engl J Med. 2007;357(9):897–904. 137. Achard C, Courtillot C, Lahuna O, et al. Normal spermatogenesis in a man with mutant luteinizing hormone. N Engl J Med. 2009;361(19):1856–1863. 138. Basciani S, Watanabe M, Mariani S, et al. Hypogonadism in a patient with two novel mutations of the luteinizing hormone beta-­ subunit gene expressed in a compound heterozygous form. J Clin Endocrinol Metab. 2012;97(9):3031–3038. 139. Potorac I, Trehan A, Szymanska K, et al. Compound heterozygous mutations in the luteinizing hormone receptor signal peptide causing 46,XY disorder of sex development. Eur J Endocrinol. 2019;181(2):K11–K20. 140. Song JW, Hwang HJ, Lee CM, et al. Hypogonadotrophic hypogonadism due to a mutation in the luteinizing hormone beta-­subunit gene. Korean J Intern Med. 2018;33(3):638–641. 141. Yang X, Ochin H, Shu L, et al. Homozygous nonsense mutation Trp28X in the LHB gene causes male hypogonadism. J Assist Reprod Genet. 2018;35(5):913–919. 142. Nagirnaja L, Venclovas C, Rull K, et al. Structural and functional analysis of rare missense mutations in human chorionic gonadotrophin beta-­subunit. Mol Hum Reprod. 2012;18(8):379–390. 143. Matthews C, Chatterjee VK. Isolated deficiency of follicle-­ stimulating hormone re-­revisited. N Engl J Med. 1997;337(9):642. 144. Matthews CH, Borgato S, Beckpeccoz P, et al. Primary amenorrhea and infertility due to a mutation in the beta-­subunit of follicle-­ stimulating-­hormone. Nat Genet. 1993;5(1):83–86. 145. Phillip M, Arbelle JE, Segev Y, Parvari R. Male hypogonadism due to a mutation in the gene for the beta-­subunit of follicle-­stimulating hormone. N Engl J Med. 1998;338(24):1729–1732. 146. Layman LC, Lee EJ, Peak DB, et al. Delayed puberty and hypogonadism caused by mutations in the follicle-­stimulating hormone beta-­subunit gene. N Engl J Med. 1997;337(9):607–611. 147. Berger K, Souza H, Brito VN, d’Alva CB, Mendonca BB, Latronico AC. Clinical and hormonal features of selective follicle-­stimulating hormone (FSH) deficiency due to FSH beta-­subunit gene mutations in both sexes. Fertil Steril. 2005;83(2):466–470. 148. Misgar RA, Wani AI, Bankura B, Bashir MI, Roy A, Das M. FSH beta-­subunit mutations in two sisters: the first report from the Indian sub-­ continent and review of previous cases. Gynecol Endocrinol. 2019;35(4):290–293. 149. Zheng J, Mao J, Cui M, et al. Novel FSHbeta mutation in a male patient with isolated FSH deficiency and infertility. Eur J Med Genet. 2017;60(6):335–339. 150. Kottler ML, Chou YY, Chabre O, et al. A new FSHbeta mutation in a 29-­ year-­ old woman with primary amenorrhea and isolated FSH deficiency: functional characterization and ovarian response to human recombinant FSH. Eur J Endocrinol. 2010;162(3):633–641. 151. Lindstedt G, Nystrom E, Matthews C, Ernest I, Janson PO, Chatterjee K. Follitropin (FSH) deficiency in an infertile male due to FSHbeta gene mutation. A syndrome of normal puberty and virilization but underdeveloped testicles with azoospermia, low FSH but high lutropin and normal serum testosterone concentrations. Clin Chem Lab Med. 1998;36(8):663–665. 152. Simsek E, Montenegro LR, Binay C, Demiral M, Acikalin MF, Latronico AC. Clinical and hormonal features of a male adolescent with congenital isolated follicle-­ stimulating hormone deficiency. Horm Res Paediatr. 2016;85(3):207–212. 153. Keir G, Winchester BG, Clayton P. Carbohydrate-­deficient glycoprotein syndromes: inborn errors of protein glycosylation. Ann Clin Biochem. 1999;36:20–36. 154. Manna PR, Joshi L, Reinhold VN, et al. Synthesis, purification and structural and functional characterization of recombinant form of a common genetic variant of human luteinizing hormone. Hum Mol Genet. 2002;11(3):301–315. 155. Wide L, Eriksson K, Sluss PM, Hall JE. The common genetic variant of luteinizing hormone has a longer serum half-­life than the wild type in heterozygous women. J Clin Endocrinol Metab. 2010;95(1):383–389. 156. Du JW, Xu KY, Fang L the luteinizing hormone beta subunit and female infertility. Rep. 2012;5(2):473–476.

157. Punab AM, Grigorova M, Punab M, et al. ‘Carriers of variant luteinizing hormone (V-­LH) among 1593 Baltic men have significantly higher serum LH’. ‘Andrology. 2015;3(3):512–519. 158. Ramanujam LN, Liao WX, Roy AC, Ng SC. Association of molecular variants of luteinizing hormone with male infertility. Hum Reprod. 2000;15(4):925–928. 159. Takahashi K, Kurioka H, Ozaki T, et al. Increased prevalence of luteinizing hormone beta-­ subunit variant in Japanese infertility patients. Hum Reprod. 1998;13(12):3338–3344. 160. Liu N, Ma Y, Wang S, et al. Association of the genetic variants of luteinizing hormone, luteinizing hormone receptor and polycystic ovary syndrome. Reprod Biol Endocrinol. 2012;10:36. 161. Kim NK, Nam YS, Ko JJ, Chung HM, Chung KW, Cha KY. The luteinizing hormone beta-­subunit exon 3 (Gly102Ser) gene mutation is rare in Korean women with endometriosis and polycystic ovary syndrome. Fertil Steril. 2001;75(6):1238–1239. 162. Miller-­Lindholm AK, Bedows E, Bartels CF, Ramey J, Maclin V, Ruddon RW. A naturally occurring genetic variant in the human chorionic gonadotropin-­ beta gene 5 is assembly inefficient. Endocrinology. 1999;140(8):3496–3506. 163. Jiang M, Savontaus ML, Simonsen H, et al. Absence of the genetic variant Val(79)Met in human chorionic gonadotropin-­ beta gene 5 in five European populations. Mol Hum Reprod. 2004;10(10):763–766. 164. Rull K, Nagirnaja L, Ulander VM, et al. Chorionic gonadotropin beta-­ gene variants are associated with recurrent miscarriage in two European populations. J Clin Endocrinol Metab. 2008;93(12):4697–4706. 165. Rull K, Christiansen OB, Nagirnaja L, Steffensen R, Margus T, Laan M. A modest but significant effect of CGB5 gene promoter polymorphisms in modulating the risk of recurrent miscarriage. Fertil Steril. 2013;99(7):1930–1936.e6. 166. Casarini L, Santi D, Marino M. Impact of gene polymorphisms of gonadotropins and their receptors on human reproductive success. Reproduction. 2015;150(6):R175–R184. 167. Grigorova M, Punab M, Ausmees K, Laan M. FSHB promoter polymorphism within evolutionary conserved element is associated with serum FSH level in men. Hum Reprod. 2008;23(9):2160–2166. 168. Ferlin A, Vinanzi C, Selice R, Garolla A, Frigo AC, Foresta C. Toward a pharmacogenetic approach to male infertility: polymorphism of follicle-­stimulating hormone beta-­subunit promoter. Fertil Steril. 2011;96(6):1344–1349.e2. 169. Grigorova M, Punab M, Poolamets O, et al. Increased Prevalance of the −211 T allele of follicle stimulating hormone (FSH) beta subunit promoter polymorphism and lower serum FSH in infertile men. J Clin Endocrinol Metab. 2010;95(1):100–108. 170. Tuttelmann F, Laan M, Grigorova M, Punab M, Sober S, Gromoll J. Combined effects of the variants FSHB −211 G>T and FSHR 2039 A>G on male reproductive parameters. J Clin Endocrinol Metab. 2012;97(10):3639–3647. 171. Rull K, Grigorova M, Ehrenberg A, et al. FSHB −211 G>T is a major genetic modulator of reproductive physiology and health in childbearing age women. Hum Reprod. 2018;33(5):954–966. 172. Benson CA, Kurz TL, Thackray VG. A human FSHB promoter SNP associated with low FSH levels in men impairs LHX3 binding and basal FSHB transcription. Endocrinology. 2013;154(9):3016–3021. 173. Day FR, Hinds DA, Tung JY, et al. Causal mechanisms and balancing selection inferred from genetic associations with polycystic ovary syndrome. Nat Commun. 2015;6:8464. 174. Hayes MG, Urbanek M, Ehrmann DA, et al. Genome-­wide association of polycystic ovary syndrome implicates alterations in gonadotropin secretion in European ancestry populations. Nat Commun. 2015;6:7502. 175. Bohaczuk SC, Thackray VG, Shen J, Skowronska-­Krawczyk D, Mellon PL. FSHB transcription is regulated by a novel 5’ distal enhancer with a fertility-­ associated single nucleotide polymorphism. Endocrinology. 2021;162(1):1–20. 176. Roa J, Navarro VM, Tena-­Sempere M. Kisspeptins in reproductive biology: consensus knowledge and recent developments. Biol Reprod. 2011;85(4):650–660. 177. Skorupskaite K, George JT, Anderson RA. The kisspeptin-­GnRH pathway in human reproductive health and disease. Hum Reprod Update. 2014;20(4):485–500. . Mechanisms for pulsatile regulation of the gonadotropin subunit genes by GNRH1. Biol Reprod. 2006;74(6):993–998.

CHAPTER 2  The Gonadotropin Hormones and Their Receptors 179. Thompson IR, Kaiser UB. GnRH pulse frequency-­ dependent differential regulation of LH and FSH gene expression. Mol Cell Endocrinol. 2014;385(1–2):28–35. 180. Garcia-­Galiano D, Pinilla L, Tena-­Sempere M. Sex steroids and the control of the Kiss1 system: developmental roles and major regulatory actions. J Neuroendocrinol. 2012;24(1):22–33. 181. Ruohonen ST, Poutanen M, Tena-­Sempere M. Role of kisspeptins in the control of the hypothalamic-­pituitary-­ovarian axis: old dogmas and new challenges. Fertil Steril. 2020;114(3):465–474. 182. Alarid ET, Holley S, Hayakawa M, Mellon PL. Discrete stages of anterior pituitary differentiation recapitulated in immortalized cell lines. Mol Cell Endocrinol. 1998;140(1–2):25–30. 183. Alarid ET, Windle JJ, Whyte DB, Mellon PL. Immortalization of pituitary cells at discrete stages of development by directed oncogenesis in transgenic mice. Development. 1996;122(10):3319–3329. 184. Haisenleder DJ, Ferris HA, Shupnik MA. The calcium component of gonadotropin-­ releasing hormone-­ stimulated luteinizing hormone subunit gene transcription is mediated by calcium/calmodulin-­dependent protein kinase type II. Endocrinology. 2003;144(6):2409–2416. 185. Thackray VG, Mellon PL, Coss D. Hormones in synergy: regulation of the pituitary gonadotropin genes. Mol Cell Endocrinol. 2010;314(2):192–203. 186. Liu F, Austin DA, Mellon PL, Olefsky JM, Webster NJ. GnRH activates ERK1/2 leading to the induction of c-­fos and LHbeta protein expression in LbetaT2 cells. Mol Endocrinol. 2002;16(3):419–434. 187. Roberson MS, Misra-­Press A, Laurance ME, Stork PJ, Maurer RA. A role for mitogen-­activated protein kinase in mediating activation of the glycoprotein hormone alpha-­subunit promoter by gonadotropin-­ releasing hormone. Mol Cell Biol. 1995;15(7):3531–3539. 188. Coss D. Regulation of reproduction via tight control of gonadotropin hormone levels. Mol Cell Endocrinol. 2018;463:116–130. 189. Stamatiades GA, Kaiser UB. Gonadotropin regulation by pulsatile GnRH: signaling and gene expression. Mol Cell Endocrinol. 2018;463:131–141. 190. Kanasaki H, Bedecarrats GY, Kam KY, Xu S, Kaiser UB. Gonadotropin-­releasing hormone pulse frequency-­dependent activation of extracellular signal-­regulated kinase pathways in perifused LbetaT2 cells. Endocrinology. 2005;146(12):5503–5513. 191. Thompson IR, Ciccone NA, Xu S, Zaytseva S, Carroll RS, Kaiser UB. GnRH pulse frequency-­ dependent stimulation of FSHbeta transcription is mediated via activation of PKA and CREB. Mol Endocrinol. 2013;27(4):606–618. 192. Thompson IR, Ciccone NA, Zhou Q, et al. GnRH pulse frequency control of fshb gene expression is mediated via ERK1/2 regulation of ICER. Mol Endocrinol. 2016;30(3):348–360. 193. Bernard DJ, Tran S. Mechanisms of activin-­stimulated FSH synthesis: the story of a pig and a FOX. Biol Reprod. 2013;88(3):78. 194. Das N, Kumar TR. Molecular regulation of follicle-­stimulating hormone synthesis, secretion and action. J Mol Endocrinol. 2018;60(3):R131–R155. 195. Stamatiades GA, Carroll RS, Kaiser UB. GnRH-­A key regulator of FSH. Endocrinology. 2019;160(1):57–67. 196. Thackray VG. Fox tales: regulation of gonadotropin gene expression by forkhead transcription factors. Mol Cell Endocrinol. 2014;385(1–2):62–70. 197. Salisbury TB, Binder AK, Nilson JH. Welcoming beta-­catenin to the gonadotropin-­ releasing hormone transcriptional network in gonadotropes. Mol Endocrinol. 2008;22(6):1295–1303. 198. Wang Q, Chikina M, Zaslavsky E, Pincas H, Sealfon SC. beta-­ catenin regulates GnRH-­induced FSHbeta gene expression. Mol Endocrinol. 2013;27(2):224–237. 199. Boerboom D, Kumar V, Boyer A, et al. Beta-­ catenin stabilization in gonadotropes impairs FSH synthesis in male mice in vivo. Endocrinology. 2015;156(1):323–333. 200. Arriola DJ, Mayo SL, Skarra DV, Benson CA, Thackray VG. FOXO1 transcription factor inhibits luteinizing hormone beta gene expression in pituitary gonadotrope cells. J Biol Chem. 2012;287(40):33424–33435. 201. Skarra DV, Arriola DJ, Benson CA, Thackray VG. Forkhead box O1 is a repressor of basal and GnRH-­induced Fshb transcription in gonadotropes. Mol Endocrinol. 2013;27(11):1825–1839. 202. Weiss J, Guendner MJ, Halvorson LM, Jameson JL. Transcriptional activation of the follicle-­ stimulating hormone beta-­ subunit gene by activin. 1885–1891.

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203. Schang G, Ongaro L, Schultz H, et al. Murine FSH production depends on the activin type II receptors ACVR2A and ACVR2B. Endocrinology. 2020;161(7):1–17. 204. Corpuz PS, Lindaman LL, Mellon PL, Coss D. FoxL2 is required for activin induction of the mouse and human follicle-­stimulating hormone beta-­subunit genes. Mol Endocrinol. 2010;24(5):1037–1051. 205. Justice NJ, Blount AL, Pelosi E, Schlessinger D, Vale W, Bilezikjian LM. Impaired FSHbeta expression in the pituitaries of Foxl2 mutant animals. Mol Endocrinol. 2011;25(8):1404–1415. 206. Lamba P, Fortin J, Tran S, Wang Y, Bernard DJ. A novel role for the forkhead transcription factor FOXL2 in activin A-­regulated follicle-­ stimulating hormone beta subunit transcription. Mol Endocrinol. 2009;23(7):1001–1013. 207. Tran S, Zhou X, Lafleur C, et al. Impaired fertility and FSH synthesis in gonadotrope-­specific Foxl2 knockout mice. Mol Endocrinol. 2013;27(3):407–421. 208. Bernard DJ, Fortin J, Wang Y, Lamba P. Mechanisms of FSH synthesis: what we know, what we don’t, and why you should care. Fertil Steril. 2010;93(8):2465–2485. 209. Coss D, Mellon PL, Thackray VG. A FoxL in the Smad house: activin regulation of FSH. Trends Endocrinol Metab. 2010;21(9):562–568. 210. Fortin J, Boehm U, Weinstein MB, Graff JM, Bernard DJ. Follicle-­ stimulating hormone synthesis and fertility are intact in mice lacking SMAD3 DNA binding activity and SMAD2 in gonadotrope cells. FASEB J. 2014;28(3):1474–1485. 211. Ongaro L, Schang G, Zhou Z, et al. Human follicle-­stimulating hormone ss subunit expression depends on FOXL2 and SMAD4. Endocrinology. 2020;161(5):1–12. 212. Roybal LL, Hambarchyan A, Meadows JD, et al. Roles of binding elements, FOXL2 domains, and interactions with cJUN and SMADs in regulation of FSHbeta. Mol Endocrinol. 2014;28(10):1640–1655. 213. Tran S, Lamba P, Wang Y, Bernard DJ. SMADs and FOXL2 synergistically regulate murine FSHbeta transcription via a conserved proximal promoter element. Mol Endocrinol. 2011;25(7):1170–1183. 214. Ruf-­Zamojski F, Fribourg M, Ge Y, et al. Regulatory architecture of the LbetaT2 gonadotrope cell underlying the response to gonadotropin-­releasing hormone. Front Endocrinol. 2018;9:34. 215. Xie H, Hoffmann HM, Iyer AK, et al. Chromatin status and transcription factor binding to gonadotropin promoters in gonadotrope cell lines. Reprod Biol Endocrinol. 2017;15(1):86. 216. Ghosh D, Sachdev S, Hannick M, Roberts R. Coordinate regulation of basal and cyclic 5’-­adenosine monophosphate (cAMP)-­activated expression of human chorionic gonadotropin-­alpha by Ets-­2 and cAMP-­ responsive element binding protein. J Mol Endocrinol. 2005;19:1049–1066. 217. LiCalsi C, Christophe S, Steger DJ, Buescher M, Fischer W, Mellon PL. AP-­ 2 family members regulate basal and cAMP-­ induced expression of human chorionic gonadotropin. Nucleic Acids Res. 2000;28(4):1036–1043. 218. Jameson JL, Hollenberg AN. Regulation of chorionic-­gonadotropin gene-­expression. Endocr Rev. 1993;14(2):203–221. 219. Johnson W, Jameson JL. AP-­2 (activating protein 2) and Sp1 (selective promoter factor 1) regulatory elements play distinct roles in the control of basal activity and cyclic adenosine 3’,5’-­monophosphate responsiveness of the human chorionic gonadotropin-­ beta promoter. Mol Endocrinol. 1999;13(11):1963–1975. 220. Steger DJ, Buscher M, Hecht JH, Mellon PL. Coordinate control of the alpha-­subunit and beta-­subunit genes of human chorionic-­ gonadotropin by trophoblast specific element-­binding protein. J Mol Endocrinol. 1993;7(12):1579–1588. 221. Liu L, Roberts RM. Silencing of the gene for the beta subunit of human chorionic gonadotropin by the embryonic transcription factor Oct-­3/4. J Biol Chem. 1996;271(28):16683–16689. 222. Glodek A, Kubiczak MJ, Walkowiak GP, Nowak-­ Markwitz E, Jankowska A. Methylation status of human chorionic gonadotropin beta subunit promoter and TFAP2A expression as factors regulating CGB gene expression in placenta. Fertil Steril. 2014;102(4):1175–1182.e8. 223. Campo S, Andreone L, Ambao V, Urrutia M, Calandra RS, Rulli SB. Hormonal regulation of follicle-­stimulating hormone glycosylation in males. Front Endocrinol. 2019;10:17. 224. Birken S, Kovalevskaya G, O’Connor J. Immunochemical measurement of early pregnancy isoforms of HCG: potential applications to fertility research, prenatal diagnosis, and cancer. Arch Med Res.

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225. Kelly LS, Birken S, Puett D. Determination of hyperglycosylated human chorionic gonadotropin produced by malignant gestational trophoblastic neoplasias and male germ cell tumors using a lectin-­based immunoassay and surface plasmon resonance. Mol Cell Endocrinol. 2007;260:33–39. 226. Kovalevskaya G, Genbacev O, Fisher SJ, Caceres E, O’Connor JF. Trophoblast origin of hCG isoforms: cytotrophoblasts are the primary source of choriocarcinoma-­like hCG. Mol Cell Endocrinol. 2002;194(1–2):147–155. 227. Wide L, Lee JY, Rasmussen C. A change in the isoforms of human chorionic gonadotropin occurs around the 13th week of gestation. J Clin Endocrinol Metab. 1994;78(6):1419–1423. 228. Birken S, McChesney R, Yershova O, et al. Patterns of LH beta cf among women in health and disease. Mol Cell Endocrinol. 2007;260:172–182. 229. Campo S, Ambao V, Creus S, et al. Carbohydrate complexity and proportions of serum FSH isoforms in the male: lectin-­based studies. Mol Cell Endocrinol. 2007;260:197–204. 230. Loreti N, Fresno C, Barrera D, et al. The glycan structure in recombinant human FSH affects endocrine activity and global gene expression in human granulosa cells. Mol Cell Endocrinol. 2013;366(1):68–80. 231. Cole LA. Biological functions of hCG and hCG-­related molecules. Reprod Biol Endocrinol. 2010;8:102. 232. Elliott MM, Kardana A, Lustbader JW, Cole LA. Carbohydrate and peptide structure of the alpha-­and beta-­subunits of human chorionic gonadotropin from normal and aberrant pregnancy and choriocarcinoma. Endocrine. 1997;7(1):15–32. 233. Kovalevskaya G, Birken S, Kakuma T, et al. Differential expression of human chorionic gonadotropin (hCG) glycosylation isoforms in failing and continuing pregnancies: preliminary characterization of the hyperglycosylated hCG epitope. J Endocrinol. 2002;172(3):497–506. 234. Birken S, Yershova O, Myers RV, Bernard MP, Moyle W. Analysis of human choriogonadotropin core 2 o-­glycan isoforms. Mol Cell Endocrinol. 2003;204(1–2):21–30. 235. Birken S. Specific measurement of o-­linked core 2 sugar-­containing isoforms of hyperglycosylated human chorionic gonadotropin by antibody b152. Tumour Biol. 2005;26(3):131–141. 236. Cole LA, Shahabi S, Oz UA, Bahado-­Singh RO, Mahoney MJ. Hyperglycosylated human chorionic gonadotropin (invasive trophoblast antigen) immunoassay: a new basis for gestational Down syndrome screening. Clin Chem. 1999;45(12):2109–2119. 237. Cole LA, Khanlian SA, Riley JM, Butler SA. Hyperglycosylated hCG in gestational implantation and in choriocarcinoma and testicular germ cell malignancy tumorigenesis. J Reprod Med. 2006;51(11):919–928. 238. Guibourdenche J, Handschuh K, Tsatsaris V, et al. Hyperglycosylated hCG is a marker of early human trophoblast invasion. J Clin Endocrinol Metab. 2010;95(10):E240–E244. 239. Handschuh K, Guibourdenche J, Tsatsaris V, et al. Human chorionic gonadotropin produced by the invasive trophoblast but not the villous trophoblast promotes cell invasion and is down-­regulated by peroxisome proliferator-­activated receptor-­gamma. Endocrinology. 2007;148(10):5011–5019. 240. Kovalevskaya G, Kakuma T, Schlatterer J, O’Connor JF. Hyperglycosylated HCG expression in pregnancy: cellular origin and clinical applications. Mol Cell Endocrinol. 2007;260:237–243. 241. de Medeiros SF, Norman RJ. Human choriogonadotrophin protein core and sugar branches heterogeneity: basic and clinical insights. Hum Reprod Update. 2009;15(1):69–95. 242. Davis JS, Kumar TR, May JV, Bousfield GR. Naturally occurring follicle-­ stimulating hormone glycosylation variants. J Glycomics Lipidomics. 2014;4(1):e117. 243. Walton WJ, Nguyen VT, Butnev VY, Singh V, Moore WT, Bousfield GR. Characterization of human FSH isoforms reveals a nonglycosylated beta-­subunit in addition to the conventional glycosylated beta-­ subunit. J Clin Endocrinol Metab. 2001;86(8):3675–3685. 244. Wang H, Butnev V, Bousfield GR, Kumar TR. A human FSHB transgene encoding the double N-­glycosylation mutant (Asn(7Delta) Asn(24Delta)) FSHbeta subunit fails to rescue Fshb null mice. Mol Cell Endocrinol. 2016;426:113–124. 245. Bousfield GR, Butnev VY, Butnev VY, Hiromasa Y, Harvey DJ, May JV. Hypo-­glycosylated human follicle-­stimulating hormone (hFSH(21/18)) is much more active in hFSH (hFSH(24)).

246. Lloyd JM, Childs GV. Differential storage and release of luteinizing hormone and follicle-­ releasing hormone from individual gonadotropes separated by centrifugal elutriation. Endocrinology. 1988;122(4):1282–1290. 247. Jablonka-­Shariff A, Boime I. Luteinizing hormone and follicle-­ stimulating hormone exhibit different secretion patterns from cultured Madin-­Darby canine kidney cells. Biol Reprod. 2004;70(3):649–655. 248. McNeilly AS, Crawford JL, Taragnat C, Nicol L, McNeilly JR. The differential secretion of FSH and LH: regulation through genes, feedback and packaging. Reprod Suppl. 2003;61:463–476. 249. Muyan M, Ryzmkiewicz DM, Boime I. Secretion of lutropin and follitropin from transfected GH3 cells: evidence for separate secretory pathways. Mol Endocrinol. 1994;8(12):1789–1797. 250. Farnworth PG. Gonadotrophin secretion revisited. How many ways can FSH leave a gonadotroph? J Endocrinol. 1995;145(3):387–395. 251. Jablonka-­Shariff A, Pearl CA, Comstock A, Boime I. A carboxyl-­ terminal sequence in the lutropin beta subunit contributes to the sorting of lutropin to the regulated pathway. J Biol Chem. 2008;283(17):11485–11492. 252. Pearl CA, Jablonka-­ Shariff A, Boime I. Rerouting of a follicle-­ stimulating hormone analog to the regulated secretory pathway. Endocrinology. 2010;151(1):388–393. 253. Jablonka-­Shariff A, Boime I. A novel carboxyl-­ terminal heptapeptide initiates the regulated secretion of LH from unique sub-­ domains of the ER. PLoS One. 2013;8(5):e65002. 254. Wang H, Larson M, Jablonka-­Shariff A, et al. Redirecting intracellular trafficking and the secretion pattern of FSH dramatically enhances ovarian function in mice. Proc Natl Acad Sci U S A. 2014;111(15):5735–5740. 255. Pearl CA, Boime I. Sulfation of LH does not affect intracellular trafficking. Mol Cell Endocrinol. 2009;309(1–2):76–81. 256. Muyan M, Boime I. Secretion of chorionic gonadotropin from human trophoblasts. Placenta. 1997;18(4):237–241. 257. Jablonka-­Shariff A, Boime I. Secretory trafficking signal encoded in the carboxyl-­terminal region of the CGbeta-­subunit. Mol Endocrinol. 2009;23(3):316–323. 258. Jablonka-­Shariff A, Garcia-­ Campayo V, Boime I. Evolution of lutropin to chorionic gonadotropin generates a specific routing signal for apical release in vivo. J Biol Chem. 2002;277(2):879–882. 259. Birken S, Maydelman Y, Gawinowicz MA, Pound A, Liu YP, Hartree AS. Isolation and characterization of human pituitary chorionic gonadotropin. Endocrinology. 1996;137(4):1402–1411. 260. Zenzmaier C, Gerth R, Gruschwitz M, Lindner H, Plas E, Berger P. Decreased levels of genuine large free hCG alpha in men presenting with abnormal semen analysis. Reprod Biol Endocrinol. 2011;9:114. 261. Cole LA. HCG variants, the growth factors which drive human malignancies. Am J Cancer Res. 2012;2(1):22–35. 262. Mitchell H, Seckl MJ. Discrepancies between commercially available immunoassays in the detection of tumour-­derived hCG. Mol Cell Endocrinol. 2007;260:310–313. 263. Iles RK. Ectopic hCG beta expression by epithelial cancer: malignant behaviour, metastasis and inhibition of tumor cell apoptosis. Mol Cell Endocrinol. 2007;260:264–270. 264. Iles RK, Delves PJ, Butler SA. Does hCG or hCGbeta play a role in cancer cell biology? Mol Cell Endocrinol. 2010;329(1–2):62–70. 265. Kölbl AC, Schlenk K, Behrendt N, Andergassen U. The importance of hCG in human endometrial adenocarcinoma and breast cancer. Int J Biol Markers. 2018;33(1):33–39. 266. Lempiainen A, Hotakainen K, Blomqvist C, Alfthan H, Stenman UH. Hyperglycosylated human chorionic gonadotropin in serum of testicular cancer patients. Clin Chem. 2012;58(7):1123–1129. 267. Lenhard M, Tsvilina A, Schumacher L, et al. Human chorionic gonadotropin and its relation to grade, stage and patient survival in ovarian cancer. BMC Cancer. 2012;12:2. 268. Sliwa A, Kubiczak M, Szczerba A, et al. Regulation of human chorionic gonadotropin beta subunit expression in ovarian cancer. BMC Cancer. 2019;19(1):746. 269. Bieche I, Lazar V, Nogues C, et al. Prognostic value of chorionic gonadotropin beta gene transcripts in human breast carcinoma. Clin Cancer Res. 1998;4(3):671–676. 270. Casadesus G, Milliken EL, Webber KM, et al. Increases in luteinizing hormone are associated with declines in cognitive performance. Mol Cell Endocrinol. 2007;269(1–2):107–111. al. Luteinizing hormone modulates cognition and amyloid-deposition in Alzheimer

CHAPTER 2  The Gonadotropin Hormones and Their Receptors APP transgenic mice. Biochim Biophys Acta, Mol Basis Dis. 2006;1762(4):447–452. 272. Webber KM, Casadesus G, Atwood CS, Bowen RL, Perry G, Smith MA. Gonadotropins: a cohesive gender-­based etiology of Alzheimer disease. Mol Cell Endocrinol. 2007;260:271–275. 273. Verdile G, Yeap BB, Clarnette RM, et al. Luteinizing hormone levels are positively correlated with plasma amyloid-­beta protein levels in elderly men. J Alzheimers Dis. 2008;14(2):201–208. 274. Bhatta S, Blair JA, Casadesus G. Luteinizing hormone involvement in aging female cognition: not all is estrogen loss. Front Endocrinol. 2018;9:544. 275. Blair JA, Bhatta S, McGee H, Casadesus G. Luteinizing hormone: evidence for direct action in the CNS. Horm Behav. 2015;76:57–62. 276. Rao CV. Involvement of luteinizing hormone in Alzheimer disease development in elderly women. Reprod Sci. 2017;24(3):355–368. 277. Verdile G, Asih PR, Barron AM, Wahjoepramono EJ, Ittner LM, Martins RN. The impact of luteinizing hormone and testosterone on beta amyloid (Abeta) accumulation: animal and human clinical studies. Horm Behav. 2015;76:81–90. 278. Bowen RL, Verdile G, Liu T, et al. Luteinizing hormone, a reproductive regulator that modulates the processing of amyloid-­ beta precursor protein and amyloid-­beta deposition. J Biol Chem. 2004;279(19):20539–20545. 279. Saberi S, Du YP, Christie M, Goldsbury C. Human chorionic gonadotropin increases beta-­cleavage of amyloid precursor protein in SH-­SY5Y cells. Cell Mol Neurobiol. 2013;33(6):747–751. 280. Lin J, Li X, Yuan F, et al. Genetic ablation of luteinizing hormone receptor improves the amyloid pathology in a mouse model of Alzheimer disease. J Neuropathol Exp Neurol. 2010;69(3):253–261. 281. Blair JA, Bhatta S, Casadesus G. CNS luteinizing hormone receptor activation rescues ovariectomy-­related loss of spatial memory and neuronal plasticity. Neurobiol Aging. 2019;78:111–120. 282. Pandian R, Cole LA, Palomaki GE. Second-­ trimester maternal serum invasive trophoblast antigen: a marker for down syndrome screening. Clin Chem. 2004;50(8):1433–1435. 283. Bahado-­Singh RO, Oz U, Shahabi S, Mahoney MJ, Baumgarten A, Cole LA. Comparison of urinary hyperglycosylated human chorionic gonadotropin concentration with the serum triple screen for Down syndrome detection in high-­risk pregnancies. Am J Obstet Gynecol. 2000;183(5):1114–1118. 284. Sturgeon CM, Ellis AR. Standardization of FSH, LH and hCG—current position and future prospects. Mol Cell Endocrinol. 2007;260:301–309. 285. Dias JA, Ulloa-­Aguirre A. New human follitropin preparations: how glycan structural differences may affect biochemical and biological function and clinical effect. Front Endocrinol. 2021:1212:636038. 286. Bousfield GR, Harvey DJ. Follicle-­stimulating hormone glycobiology. Endocrinology. 2019;160:1515–1535. 287. Wide L, Eriksson K. Low-­glycosylated forms of both FSH and LH play major roles in the natural ovarian stimulation. Ups J Med Sci. 2018;123(2):100–108. 288. Szkudlinski MW. New frontier in glycoprotein hormones and their receptors structure-­function. Front Endocrinol. 2015;6:155. 289. Perlman S, van den Hazel B, Christiansen J, et al. Glycosylation of an N-­terminal extension prolongs the half-­life and increases the in vivo activity of follicle stimulating hormone. J Clin Endocrinol Metab. 2003;88(7):3227–3235. 290. Fares FA, Suganuma N, Nishimori K, Boime I. Design of a longacting follitropin agonist by fusing the C-terminal sequence of the chorionic gonadotropin beta subunit to the follotropin beta subunit. Proc Natl Acad Sci USA. 1992;89(10):4304–4308. 291. Fares FA, Suganuma N, Nishimori K, LaPolt PS, Hsueh AJ, Boime I. Design of a long-­acting follitropin agonist by fusing the C-­terminal sequence of the chorionic gonadotropin beta subunit to the follitropin beta subunit. Proc Natl Acad Sci U S A. 1992;89(10):4304–4308. 292. Fauser BC, Mannaerts BM, Devroey P, Leader A, Boime I, Baird DT. Advances in recombinant DNA technology: corifollitropin alfa, a hybrid molecule with sustained follicle-­stimulating activity and reduced injection frequency. Hum Reprod Update. 2009;15(3):309–321. 293. Loutradis D, Drakakis P, Vlismas A, Antsaklis A. Corifollitropin alfa, a long-­acting follicle-­stimulating hormone agonist for the treatment of infertility. Curr Opin Investig Drugs. 2009;10(4):372–380. 294. Low SC, Nunes SL, Bitonti AJ, Dumont JA. Oral and pulmonary delivery of FSHmediated transcytosis. Hum Reprod

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295. Fredriksson R, Lagerstrom MC, Lundin LG, Schioth HB. The G-­protein-­coupled receptors in the human genome form five main families. Phylogenetic analysis, paralogon groups, and fingerprints. Mol Pharmacol. 2003;63(6):1256–1272. 296. Ascoli M, Fanelli F, Segaloff DL. The lutropin/choriogonadotropin receptor. A 2002 perspective. Endocr Rev. 2002;23:141–174. 297. Ulloa-­Aguirre A, Zarinan T. The follitropin receptor: matching structure and function. Mol Pharmacol. 2016;90(5):596–608. 298. Costagliola S, Urizar E, Mendive F, Vassart G. Specificity and promiscuity of gonadotropin receptors. Reproduction. 2005;130(3):275–281. 299. Kleinau G, Krause G. Thyrotropin and homologous glycoprotein hormone receptors: structural and functional aspects of extracellular signaling mechanisms. Endocr Rev. 2009;30(2):133–151. 300. Vassart G, Pardo L, Costagliola S. A molecular dissection of the glycoprotein hormone receptors. Trends Biochem Sci. 2004;29(3):119–126. 301. Bogerd J. Ligand-­selective determinants in gonadotropin receptors. Mol Cell Endocrinol. 2007;260–262:144–152. 302. Petrie EJ, Lagaida S, Sethi A, Bathgate RA, Gooley PR. In a class of their own—RXFP1 and RXFP2 are unique members of the LGR family. Front Endocrinol. 2015;6:137. 303. Mueller S, Jaeschke H, Gunther R, Paschke R. The hinge region: an important receptor component for GPHR function. Trends Endocrinol Metab. 2009;21(2):111–122. 304. Alvarez CA, Narayan P, Huang J, Puett D. Characterization of a region of the lutropin receptor extracellular domain near transmembrane helix 1 that is important in ligand-­mediated signaling. Endocrinology. 1999;140(4):1775–1782. 305. Brüser A, Schulz A, Rothemund S, et al. The activation mechanism of glycoprotein hormone receptors with implications in the cause and therapy of endocrine diseases. J Biol Chem. 2016;291(2):508–520. 306. Troppmann B, Kleinau G, Krause G, Gromoll J. Structural and functional plasticity of the luteinizing hormone/choriogonadotrophin receptor. Hum Reprod Update. 2013;19(5):583–602. 307. Jiang X, Dreano M, Buckler DR, et al. Structural predictions for the ligand-­ binding region of glycoprotein hormone receptors and the nature of hormone-­receptor interactions. Structure. 1995;3(12):1341–1353. 308. Schmidt A, MacColl R, Lindau-­Shepard B, Buckler DR, Dias JA. Hormone-­induced conformational change of the purified soluble hormone binding domain of follitropin receptor complexed with single chain follitropin. J Biol Chem. 2001;276(26):23373–23381. 309. Agrawal G, Dighe RR. Critical involvement of the hinge region of the follicle-­stimulating hormone receptor in the activation of the receptor. J Biol Chem. 2009;284(5):2636–2647. 310. Bruysters M, Verhoef-­Post M, Themmen AP. Asp330 and Tyr331 in the C-­terminal cysteine-­rich region of the luteinizing hormone receptor are key residues in hormone-­induced receptor activation. J Biol Chem. 2008;283(38):25821–25828. 311. Costagliola S, Panneels V, Bonomi M, et al. Tyrosine sulfation is required for agonist recognition by glycoprotein hormone receptors. EMBO J. 2002;21(4):504–513. 312. Majumdar R, Dighe RR. The hinge region of human thyroid-­ stimulating hormone (TSH) receptor operates as a tunable switch between hormone binding and receptor activation. PLoS One. 2012;7(7):e40291. 313. Fan QR, Hendrickson WA. Assembly and structural characterization of an authentic complex between human follicle stimulating hormone and a hormone-­binding ectodomain of its receptor. Mol Cell Endocrinol. 2007;260–262:73–82. 314. Jiang X, Dias JA, He X. Structural biology of glycoprotein hormones and their receptors: insights to signaling. Mol Cell Endocrinol. 2014;382(1):424–451. 315. Chen CR, Chazenbalk GD, McLachlan SM, Rapoport B. Evidence that the C terminus of the A subunit suppresses thyrotropin receptor constitutive activity. Endocrinology. 2003;144(9):3821–3827. 316. Vlaeminck-­Guillem V, Ho SC, Rodien P, Vassart G, Costagliola S. Activation of the cAMP pathway by the TSH receptor involves switching of the ectodomain from a tethered inverse agonist to an agonist. Mol Endocrinol. 2002;16(4):736–746. 317. Zhang M, Tong KP, Fremont V, et al. The extracellular domain suppresses constitutive activity of the transmembrane domain of the human TSH receptor: implications for hormone-­ receptor interaction and antagonist design. Endocrinology.

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55.e8

PART I  The Fundamentals of Reproduction

318. Nakabayashi K, Kudo M, Kobilka B, Hsueh AJ. Activation of the luteinizing hormone receptor following substitution of Ser-­277 with selective hydrophobic residues in the ectodomain hinge region. J Biol Chem. 2000;275(39):30264–30271. 319. Grzesik P, Kreuchwig A, Rutz C, et al. Differences in signal activation by LH and hCG are mediated by the LH/CG receptor’s extracellular hinge region. Front Endocrinol. 2015;6:140. 320. Bonomi M, Busnelli M, Persani L, Vassart G, Costagliola S. Structural differences in the hinge region of the glycoprotein hormone receptors: evidence from the sulfated tyrosine residues. Mol Endocrinol. 2006;20(12):3351–3363. 321. Casarini L, Lispi M, Longobardi S, et al. LH and hCG action on the same receptor results in quantitatively and qualitatively different intracellular signalling. PLoS One. 2012;7(10):e46682. 322. Jonas KC, Fanelli F, Huhtaniemi IT, Hanyaloglu AC. Single molecule analysis of functionally asymmetric G protein-­coupled receptor (GPCR) oligomers reveals diverse spatial and structural assemblies. J Biol Chem. 2015;290(7):3875–3892. 323. Grzesik P, Teichmann A, Furkert J, et al. Differences between lutropin-­mediated and choriogonadotropin-­mediated receptor activation. FEBS J. 2014;281(5):1479–1492. 324. Sanders J, Chirgadze DY, Sanders P, et al. Crystal structure of the TSH receptor in complex with a thyroid-­stimulating autoantibody. Thyroid. 2007;17(5):395–410. 325. Sanders J, Miguel RN, Bolton J, et al. Molecular interactions between the TSH receptor and a Thyroid-­stimulating monoclonal autoantibody. Thyroid. 2007;17(8):699–706. 326. Sanders P, Young S, Sanders J, et al. Crystal structure of the TSH receptor (TSHR) bound to a blocking-­type TSHR autoantibody. J Mol Endocrinol. 2011;46(2):81–99. 327. Kumar P, Sait SF, Sharma A, Kumar M. Ovarian hyperstimulation syndrome. J Hum Reprod Sci. 2011;4(2):70–75. 328. De Leener A, Caltabiano G, Erkan S, et al. Identification of the first germline mutation in the extracellular domain of the follitroplin receptor responsible for spontaneous ovarian Hyperstimulation syndrome. Hum Mutat. 2008;29(1):91–98. 329. Dupakuntla M, Mahale SD. Accessibility of the extracellular loops of follicle stimulating hormone receptor and their role in hormone-­receptor interaction. Mol Cell Endocrinol. 2010;315(1–2): 131–137. 330. Fan QR, Hendrickson WA. Structural biology of glycoprotein hormones and their receptors. Endocrine. 2005;26(3):179–188. 331. Banerjee AA, Mahale SD. Extracellular loop 3 substitutions K589N and A590S in FSH receptor increase FSH-­ induced receptor internalization and along with S588T substitution exhibit impaired ERK1/2 phosphorylation. Arch Biochem Biophys. 2018;659:57–65. 332. Banerjee AA, Dupakuntla M, Pathak BR, Mahale SD. FSH receptor-­ specific residues L501 and I505 in extracellular loop 2 are essential for its function. J Mol Endocrinol. 2015;54(3):193–204. 333. Casas-­Gonzalez P, Scaglia HE, Perez-­Solis MA, et al. Normal testicular function without detectable follicle-­stimulating hormone. A novel mutation in the follicle-­stimulating hormone receptor gene leading to apparent constitutive activity and impaired agonist-­ induced desensitization and internalization. Mol Cell Endocrinol. 2012;364(1–2):71–82. 334. Ji I, Ji TH. Differential roles of exoloop 1 of the human follicle-­ stimulating hormone receptor in hormone binding and receptor activation. J Biol Chem. 1995;270(27):15970–15973. 335. Ryu K, Gilchrist RL, Tung CS, Ji I, Ji TH. High affinity hormone binding to the extracellular N-­terminal exodomain of the follicle-­ stimulating hormone receptor is critically modulated by exoloop 3. J Biol Chem. 1998;273(44):28953–28958. 336. Ryu K, Lee H, Kim S, et al. Modulation of high affinity hormone binding. Human choriogonadotropin binding to the exodomain of the receptor is influenced by exoloop 2 of the receptor. J Biol Chem. 1998;273(11):6285–6291. 337. Banerjee AA, Mahale SD. Role of the extracellular and intracellular loops of follicle-­stimulating hormone receptor in its function. Front Endocrinol. 2015;6:110. 338. Dupakuntla M, Pathak B, Roy BS, Mahale SD. Extracellular loop 2 in the FSH receptor is crucial for ligand mediated receptor activation. Mol Cell Endocrinol. 2012;362(1–2):60–68. 339. Sohn J, Ryu K, Sievert G, Jeoung M, Ji I, Ji TH. Follicle-­stimulating hormone interacts with exoloop 3 of the receptor. 2002;277(51):50165–50175.

340. Touraine P, Beau I, Gougeon A, et al. New natural inactivating mutations of the follicle-­ stimulating hormone receptor: correlations between receptor function and phenotype. Mol Endocrinol. 1999;13(11):1844–1854. 341. Kleinau G, Claus M, Jaeschke H, et al. Contacts between extracellular loop two and transmembrane helix six determine basal activity of the thyroid-­ stimulating hormone receptor. J Biol Chem. 2007;282(1):518–525. 342. Wolf S, Grunewald S. Sequence, structure and ligand binding evolution of rhodopsin-­like G protein-­coupled receptors: a crystal structure-­ based phylogenetic analysis. PLoS One. 2015;10(4):e0123533. 343. Ulloa-­Aguirre A, Reiter E, Bousfield G, Dias JA, Huhtaniemi I. Constitutive activity in gonadotropin receptors. Adv Pharmacol. 2014;70:37–80. 344. Cohen BD, Bariteau JT, Magenis LM, Dias JA. Regulation of follitropin receptor cell surface residency by the ubiquitin-­proteasome pathway. Endocrinology. 2003;144(10):4393–4402. 345. Cohen BD, Nechamen CA, Dias JA. Human follitropin receptor (FSHR) interacts with the adapter protein 14-­3-­3tau. Mol Cell Endocrinol. 2004;220(1–2):1–7. 346. Dias JA, Mahale SD, Nechamen CA, Davydenko O, Thomas RM, Ulloa-­Aguirre A. Emerging roles for the FSH receptor adapter protein APPL1 and overlap of a putative 14-­3-­3tau interaction domain with a canonical G-­protein interaction site. Mol Cell Endocrinol. 2010;329(1–2):17–25. 347. Timossi C, Maldonado D, Vizcaino A, Lindau-­Shepard B, Conn PM, Ulloa-­Aguirre A. Structural determinants in the second intracellular loop of the human follicle-­ stimulating hormone receptor are involved in G(s) protein activation. Mol Cell Endocrinol. 2002;189(1–2):157–168. 348. Timossi C, Ortiz-­Elizondo C, Pineda DB, Dias JA, Conn PM, Ulloa-­ Aguirre A. Functional significance of the BBXXB motif reversed present in the cytoplasmic domains of the human follicle-­stimulating hormone receptor. Mol Cell Endocrinol. 2004;223(1–2):17–26. 349. Schulz A, Schoneberg T, Paschke R, Schultz G, Gudermann T. Role of the third intracellular loop for the activation of gonadotropin receptors. Mol Endocrinol. 1999;13(2):181–190. 350. Scheerer P, Park JH, Hildebrand PW, et al. Crystal structure of opsin in its G-­ protein-­ interacting conformation. Nature. 2008;455(7212):497–502. 351. Rasmussen SGF, Choi HJ, Fung JJ, et al. Structure of a nanobody-­ stabilized active state of the beta(2) adrenoceptor. Nature. 2011;469(7329):175–180. 352. Rasmussen SGF, DeVree BT, Zou YZ, et al. Crystal structure of the beta(2) adrenergic receptor-­ Gs protein complex. Nature. 2011;477(7366). 549–555. 353. Venkatakrishnan AJ, Deupi X, Lebon G, Tate CG, Schertler GF, Babu MM. Molecular signatures of G-­protein-­coupled receptors. Nature. 2013;494(7436):185–194. 354. Ballesteros JA, Weinstein H. Analysis and refinement of criteria for predicting the structure and relative orientations of transmembranal helical domains. Biochem J. 1992;62(1):107–109. 355. Deupi X, Standfuss J. Structural insights into agonist-­ induced activation of G-­protein-­coupled receptors. Curr Opin Struct Biol. 2011;21(4):541–551. 356. Sansuk K, Deupi X, Torrecillas IR, et al. A structural insight into the reorientation of transmembrane domains 3 and 5 during family A G protein-­ coupled receptor activation. Mol Pharmacol. 2011;79(2):262–269. 357. Standfuss J, Edwards PC, D’Antona A, et al. The structural basis of agonist-­ induced activation in constitutively active rhodopsin. Nature. 2011;471(7340):656–660. 358. Nygaard R, Zou Y, Dror RO, et al. The dynamic process of beta(2)-­ adrenergic receptor activation. Cell. 2013;152(3):532–542. 359. Ayoub MA, Yvinec R, Crepieux P, Poupon A. Computational modeling approaches in gonadotropin signaling. Theriogenology. 2016;86(1):22–31. 360. Landomiel F, Gallay N, Jegot G, et al. Biased signalling in follicle stimulating hormone action. Mol Cell Endocrinol. 2014;382(1):452–459. 361. Melo-­Nava B, Casas-­Gonzalez P, Perez-­Solis MA, et al. Role of cysteine residues in the carboxyl-­terminus of the follicle-­stimulating hormone receptor in intracellular traffic and postendocytic processing. Front Cell Dev Biol. 2016;4:76. 362. Munshi UM, Clouser CL, Peegel H, Menon KM. Evidence that palmitoylation of carboxyl terminus cysteine residues of the human luteinizing hormone receptor regulates postendocytic processing.

CHAPTER 2  The Gonadotropin Hormones and Their Receptors 363. Uribe A, Zarinan T, Perez-­Solis MA, et al. Functional and structural roles of conserved cysteine residues in the carboxyl-­terminal domain of the follicle-­stimulating hormone receptor in human embryonic kidney 293 cells. Biol Reprod. 2008;78:869–882. 364. Duvernay MT, Zhou F, Wu G. A conserved motif for the transport of G protein-­coupled receptors from the endoplasmic reticulum to the cell surface. J Biol Chem. 2004;279(29):30741–30750. 365. Kara E, Crepieux P, Gauthier C, et al. A phosphorylation cluster of five serine and threonine residues in the C-­terminus of the follicle-­ stimulating hormone receptor is important for desensitization but not for beta-­ arrestin-­ mediated ERK activation. Mol Endocrinol. 2006;20(11):3014–3026. 366. Lazari MF, Liu X, Nakamura K, Benovic JL, Ascoli M. Role of G protein-­coupled receptor kinases on the agonist-­induced phosphorylation and internalization of the follitropin receptor. Mol Endocrinol. 1999;13(6):866–878. 367. Troispoux C, Guillou F, Elalouf JM, et al. Involvement of G protein-­ coupled receptor kinases and arrestins in desensitization to follicle-­ stimulating hormone action. Mol Endocrinol. 1999;13(9):1599–1614. 368. Krishnamurthy H, Kishi H, Shi M, et al. Postendocytotic trafficking of the follicle-­stimulating hormone (FSH)-­FSH receptor complex. Mol Endocrinol. 2003;17(11):2162–2176. 369. Galet C, Hirakawa T, Ascoli M. The postendocytotic trafficking of the human lutropin receptor is mediated by a transferable motif consisting of the C-­terminal cysteine and an upstream leucine. Mol Endocrinol. 2004;18(2):434–446. 370. Galet C, Min L, Narayanan R, Kishi M, Weigel NL, Ascoli M. Identification of a transferable two-­amino-­acid motif (GT) present in the C-­terminal tail of the human lutropin receptor that redirects internalized G protein-­coupled receptors from a degradation to a recycling pathway. Mol Endocrinol. 2003;17(3):411–422. 371. Chauvigne F, Tingaud-­Sequeira A, Agulleiro MJ, et al. Functional and evolutionary analysis of flatfish gonadotropin receptors reveals cladal-­and lineage-­ level divergence of the teleost glycoprotein receptor family. Biol Reprod. 2010;82(6):1088–1102. 372. Dufau ML, Tsai-­Morris CH. The luteinizing hormone receptor. In: Payne AH, Hardy MP, eds. The Leydig Cell in Health and Disease. Humana Press, Totowa NJ; 2007:227–252. 373. George JW, Dille EA, Heckert LL. Current concepts of follicle-­ stimulating hormone receptor gene regulation. Biol Reprod. 2011;84(1):7–17. 374. Hermann BP, Heckert LL. Silencing of Fshr occurs through a conserved, hypersensitive site in the first intron. Mol Endocrinol. 2005;19(8):2112–2131. 375. Hermann BP, Heckert LL. Transcriptional regulation of the FSH receptor: new perspectives. Mol Cell Endocrinol. 2007;260–262:100–108. 376. Hermann BP, Hornbaker K, Rice DA, Sawadogo M, Heckert LL. In vivo regulation of follicle-­ stimulating hormone receptor by the transcription factors upstream stimulatory factor 1 and upstream stimulatory factor 2 is cell specific. Endocrinology. 2008;149(10):5297–5306. 377. Hermann BP, Hornbaker KI, Maran RR, Heckert LL. Distal regulatory elements are required for Fshr expression, in vivo. Mol Cell Endocrinol. 2007;260–262:49–58. 378. Scobey MJ, Fix CA, Walker WH. The Id2 transcriptional repressor is induced by follicle-­stimulating hormone and cAMP. J Biol Chem. 2004;279(16):16064–16070. 379. Viswanathan P, Wood MA, Walker WH. Follicle-­stimulating hormone (FSH) transiently blocks FSH receptor transcription by increasing inhibitor of deoxyribonucleic acid binding/differentiation-­2 and decreasing upstream stimulatory factor expression in rat Sertoli cells. Endocrinology. 2009;150(8):3783–3791. 380. Wood MA, Walker WH. USF1/2 transcription factor DNA-­ binding activity is induced during rat Sertoli cell differentiation. Biol Reprod. 2009;80(1):24–33. 381. Gromoll J, Dankbar B, Gudermann T. Characterization of the 5’ flanking region of the human follicle-­stimulating hormone receptor gene. Mol Cell Endocrinol. 1994;102(1–2):93–102. 382. Levallet J, Koskimies P, Rahman N, Huhtaniemi I. The promoter of murine follicle-­stimulating hormone receptor: functional characterization and regulation by transcription factor steroidogenic factor 1. Mol Endocrinol. 2001;15(1):80–92. 383. Perez-­Solis MA, Macias H, Acosta-­MontesdeOca A, et al. Molecular cloning and functional analysis of the FSH receptor gene promoter

55.e9

from the volcano mouse (Neotomodon alstoni alstoni). Endocrine. 2010;37(1):98–105. 384. Sairam MR, Jiang LG, Yarney TA, Khan H. Alternative splicing converts the G-­protein coupled follitropin receptor gene into a growth factor type I receptor: implications for pleiotropic actions of the hormone. Mol Reprod Dev. 1997;48(4):471–479. 385. Dufau ML, Liao M, Zhang Y. Participation of signaling pathways in the derepression of luteinizing hormone receptor transcription. Mol Cell Endocrinol. 2010;314(2):221–227. 386. El-­Hefnawy T, Krawczyk Z, Nikula H, Vihera I, Huhtaniemi I. Regulation of function of the murine luteinizing hormone receptor promoter by cis-­and trans-­acting elements in mouse Leydig tumor cells. Mol Cell Endocrinol. 1996;119(2):207–217. 387. Geng Y, Tsai-­ Morris CH, Zhang Y, Dufau ML. The human luteinizing hormone receptor gene promoter: activation by Sp1 and Sp3 and inhibitory regulation. Biochem Biophys Res Commun. 1999;263(2):366–371. 388. Zhang Y, Dufau ML. Repression of the luteinizing hormone receptor gene promoter by cross talk among EAR3/COUP-­TFI, Sp1/ Sp3, and TFIIB. Mol Cell Biol. 2003;23(19):6958–6972. 389. Zhang Y, Fatima N, Dufau ML. Coordinated changes in DNA methylation and histone modifications regulate silencing/derepression of luteinizing hormone receptor gene transcription. Mol Cell Biol. 2005;25(18):7929–7939. 390. Zhang Y, Liao M, Dufau ML. Unlocking repression of the human luteinizing hormone receptor gene by trichostatin A-­induced cell-­ specific phosphatase release. J Biol Chem. 2008;283(35):24039–24046. 391. Zhang Y, Dufau ML. Dual mechanisms of regulation of transcription of luteinizing hormone receptor gene by nuclear orphan receptors and histone deacetylase complexes. J Steroid Biochem Mol Biol. 2003;85(2–5):401–414. 392. Liao M, Zhang Y, Kang JH, Dufau ML. Coactivator function of positive cofactor 4 (PC4) in Sp1-­directed luteinizing hormone receptor (LHR) gene transcription. J Biol Chem. 2011;286(9):7681–7691. 393. Liao M, Zhang Y, Dufau ML. Protein kinase Calpha-­induced derepression of the human luteinizing hormone receptor gene transcription through ERK-­mediated release of HDAC1/Sin3A repressor complex from Sp1 sites. Mol Endocrinol. 2008;22(6):1449–1463. 394. Simoni M, Gromoll J, Nieschlag E. The follicle-­stimulating hormone receptor: biochemistry, molecular biology, physiology, and pathophysiology. Endocr Rev. 1997;18(6):739–773. 395. LaPolt PS, Tilly JL, Aihara T, Nishimori K, Hsueh AJ. Gonadotropin-­induced up-­and down-­regulation of ovarian follicle-­ stimulating hormone (FSH) receptor gene expression in immature rats: effects of pregnant mare’s serum gonadotropin, human chorionic gonadotropin, and recombinant FSH. Endocrinology. 1992;130(3):1289–1295. 396. Rannikki AS, Zhang FP, Huhtaniemi IT. Ontogeny of follicle-­ stimulating hormone receptor gene expression in the rat testis and ovary. Mol Cell Endocrinol. 1995;107(2):199–208. 397. Babu PS, Danilovich N, Sairam MR. Hormone-­ induced receptor gene splicing: enhanced expression of the growth factor type I follicle-­ stimulating hormone receptor motif in the developing mouse ovary as a new paradigm in growth regulation. Endocrinology. 2001;142(1):381–389. 398. Gromoll J, Dankbar B, Sharma RS, Nieschlag E. Molecular cloning of the testicular follicle stimulating hormone receptor of the non human primate Macaca fascicularis and identification of multiple transcripts in the testis. Biochem Biophys Res Commun. 1993;196(3):1066–1072. 399. Houde A, Lambert A, Saumande J, Silversides DW, Lussier JG. Structure of the bovine follicle-­stimulating hormone receptor complementary DNA and expression in bovine tissues. Mol Reprod Dev. 1994;39(2):127–135. 400. Khan H, Yarney TA, Sairam MR. Cloning of alternately spliced mRNA transcripts coding for variants of ovine testicular follitropin receptor lacking the G protein coupling domains. Biochem Biophys Res Commun. 1993;190(3):888–894. 401. Neubauer K, Jewgenow K, Fickel J. Follicle-­stimulating hormone receptor in felids: intra-­and interspecies variation. Theriogenology. 2006;66(6–7):1737–1742. 402. Rajapaksha WR, Robertson L, O’Shaughnessy PJ. Expression of follicle-­ stimulating hormone-­ receptor mRNA alternate transcripts in bovine granulosa cells during luteinization in vivo and . 1996;120(1):25–30.

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55.e10 PART I  The Fundamentals of Reproduction 403. Tena-­Sempere M, Manna PR, Huhtaniemi I. Molecular cloning of the mouse follicle-­stimulating hormone receptor complementary deoxyribonucleic acid: functional expression of alternatively spliced variants and receptor inactivation by a C566T transition in exon 7 of the coding sequence. Biol Reprod. 1999;60(6):1515–1527. 404. Gerasimova T, Thanasoula MN, Zattas D, Seli E, Sakkas D, Lalioti MD. Identification and in vitro characterization of follicle stimulating hormone (FSH) receptor variants associated with abnormal ovarian response to FSH. J Clin Endocrinol Metab. 2010;95(2):529–536. 405. Gromoll J, Gudermann T, Nieschlag E. Molecular cloning of a truncated isoform of the human follicle stimulating hormone receptor. Biochem Biophys Res Commun. 1992;188(3):1077–1083. 406. Karakaya C, Guzeloglu-­Kayisli O, et al. Follicle-­stimulating hormone receptor (FSHR) alternative skipping of exon 2 or 3 affects ovarian response to FSH. Mol Hum Reprod. 2014;20(7):630–643. 407. Kelton CA, Cheng SV, Nugent NP, et al. The cloning of the human follicle stimulating hormone receptor and its expression in COS-­7, CHO, and Y-­1 cells. Mol Cell Endocrinol. 1992;89(1–2):141–151. 408. Robinson LJ, Tourkova I, Wang Y, et al. FSH-­receptor isoforms and FSH-­dependent gene transcription in human monocytes and osteoclasts. Biochem Biophys Res Commun. 2010;394(1):12–17. 409. Song GJ, Park YS, Lee YS, Lee CC, Kang IS. Alternatively spliced variants of the follicle-­stimulating hormone receptor gene in the testis of infertile men. Fertil Steril. 2002;77(3):499–504. 410. Zhou J, Huang Y, Li L, et al. Identification of two novel, alternatively spliced mRNA transcripts of the human follicle-­stimulating hormone receptor. Mol Reprod Dev. 2013;80(11):916–923. 411. Sairam MR, Babu PS. The tale of follitropin receptor diversity: a recipe for fine tuning gonadal responses? Mol Cell Endocrinol. 2007;260–262:163–171. 412. Babu PS, Jiang L, Sairam AM, Touyz RM, Sairam MR. Structural features and expression of an alternatively spliced growth factor type I receptor for follitropin signaling in the developing ovary. Mol Cell Biol Res Commun. 1999;2(1):21–27. 413. Babu PS, Krishnamurthy H, Chedrese PJ, Sairam MR. Activation of extracellular-­regulated kinase pathways in ovarian granulosa cells by the novel growth factor type 1 follicle-­stimulating hormone receptor. Role in hormone signaling and cell proliferation. J Biol Chem. 2000;275(36):27615–27626. 414. Sairam MR, Jiang LG, Yarney TA, Khan H. Follitropin signal transduction: alternative splicing of the FSH receptor gene produces a dominant negative form of receptor which inhibits hormone action. Biochem Biophys Res Commun. 1996;226(3):717–722. 415. Sairam MR, Subbarayan VS. Characterization of the 5’ flanking region and potential control elements of the ovine follitropin receptor gene. Mol Reprod Dev. 1997;48(4):480–487. 416. Sullivan RR, Faris BR, Eborn D, Grieger DM, Cino-­Ozuna AG, Rozell TG. Follicular expression of follicle stimulating hormone receptor variants in the Ewe. Reprod Biol Endocrinol. 2013;11:113. 417. McFarland KC, Sprengel R, Phillips HS, et al. Lutropin-­ choriogonadotropin receptor: an unusual member of the G protein-­ coupled receptor family. Science. 1989;245(4917):494–499. 418. Minegishi T, Nakamura K, Takakura Y, et al. Cloning and sequencing of human LH/hCG receptor cDNA. Biochem Biophys Res Commun. 1990;172(3):1049–1054. 419. Bacich DJ, Rohan RM, Norman RJ, Rodgers RJ. Characterization and relative abundance of alternatively spliced luteinizing hormone receptor messenger ribonucleic acid in the ovine ovary. Endocrinology. 1994;135(2):735–744. 420. Kawate N, Okuda K. Coordinated expression of splice variants for luteinizing hormone receptor messenger RNA during the development of bovine corpora lutea. Mol Reprod Dev. 1998;51(1):66–75. 421. Saint-­Dizier M, Chopineau M, Dupont J, Daels PF, Combarnous Y. Expression and binding activity of luteinizing hormone/chorionic gonadotropin receptors in the primary corpus luteum during early pregnancy in the mare. Biol Reprod. 2003;69(5):1743–1749. 422. Igarashi S, Minegishi T, Nakamura K, et al. Functional expression of recombinant human luteinizing hormone/human choriogonadotropin receptor. Biochem Biophys Res Commun. 1994;201(1):248–256. 423. Madhra M, Gay E, Fraser HM, Duncan WC. Alternative splicing of the human luteal LH receptor during luteolysis and maternal recognition of pregnancy. Mol Hum Reprod. 2004;10(8):599–603. 424. Minegishi T, Nakamura K, Yamashita S, Omori Y. The effect of splice variant of the human luteinizing hormone (LH) receptor on the expression of gonadotropin receptor. 2007;260–262:117–125.

425. Nakamura K, Yamashita S, Omori Y, Minegishi T. A splice variant of the human luteinizing hormone (LH) receptor modulates the expression of wild-­type human LH receptor. Mol Endocrinol. 2004;18(6):1461–1470. 426. Nishimori K, Dunkel L, Hsueh AJ, Yamoto M, Nakano R. Expression of luteinizing hormone and chorionic gonadotropin receptor messenger ribonucleic acid in human corpora lutea during menstrual cycle and pregnancy. J Clin Endocrinol Metab. 1995;80(4):1444–1448. 427. Yamashita S, Nakamura K, Omori Y, Tsunekawa K, Murakami M, Minegishi T. Association of human follitropin (FSH) receptor with splicing variant of human lutropin/choriogonadotropin receptor negatively controls the expression of human FSH receptor. Mol Endocrinol. 2005;19(8):2099–2111. 428. Kossack N, Simoni M, Richter-­Unruh A, Themmen AP, Gromoll J. Mutations in a novel, cryptic exon of the luteinizing hormone/ chorionic gonadotropin receptor gene cause male pseudohermaphroditism. PLoS Med. 2008;5(4):e88. 429. Kossack N, Troppmann B, Richter-­Unruh A, Kleinau G, Gromoll J. Aberrant transcription of the LHCGR gene caused by a mutation in exon 6A leads to Leydig cell hypoplasia type II. Mol Cell Endocrinol. 2013;366(1):59–67. 430. Menon B, Franzo-­ Romain M, Damanpour S, Menon KM. Luteinizing hormone receptor mRNA down-­regulation is mediated through ERK-­dependent induction of RNA binding protein. Mol Endocrinol. 2011;25(2):282–290. 431. Gulappa T, Menon B, Menon KM. Hypusination of eukaryotic initiation factor 5A via cAMP-­PKA-­ERK1/2 pathway is required for ligand-­induced downregulation of LH receptor mRNA expression in the ovary. Mol Cell Endocrinol. 2015;413:90–95. 432. Menon B, Sinden J, Franzo-­Romain M, Botta RB, Menon KM. Regulation of LH receptor mRNA binding protein by miR-­122 in rat ovaries. Endocrinology. 2013;154(12):4826–4834. 433. Menon KM, Menon B. Structure, function and regulation of gonadotropin receptors—a perspective. Mol Cell Endocrinol. 2012;356(1–2):88–97. 434. Menon B, Gulappa T, Menon KM. miR-­122 regulates LH receptor expression by activating sterol response element binding protein in rat ovaries. Endocrinology. 2015;156(9):3370–3380. 435. Menon B, Guo X, Garcia N, Gulappa T, Menon KMJ. miR-­122 regulates LHR expression in rat granulosa cells by targeting Insig1 mRNA. Endocrinology. 2018;159(5):2075–2082. 436. Troppmann B, Kossack N, Nordhoff V, Schuring AN, Gromoll J. MicroRNA miR-­513a-­3p acts as a co-­regulator of luteinizing hormone/chorionic gonadotropin receptor gene expression in human granulosa cells. Mol Cell Endocrinol. 2014;390(1–2):65–72. 437. Kitahara Y, Nakamura K, Kogure K, Minegishi T. Role of microRNA-­ 136-­ 3p on the expression of luteinizing hormone-­ human chorionic gonadotropin receptor mRNA in rat ovaries. Biol Reprod. 2013;89(5):114. 438. Inoue K, Nakamura K, Abe K, et al. Effect of transforming growth factor beta on the expression of luteinizing hormone receptor in cultured rat granulosa cells. Biol Reprod. 2002;67(2):610–615. 439. Tano M, Minegishi T, Nakamura K, Karino S, Ibuki Y, Miyamoto K. Transcriptional and post-­transcriptional regulation of FSH receptor in rat granulosa cells by cyclic AMP and activin. J Endocrinol. 1997;153(3):465–473. 440. Du X, Li Q, Pan Z, Li Q. Androgen receptor and miRNA-­126* axis controls follicle-­stimulating hormone receptor expression in porcine ovarian granulosa cells. Reproduction. 2016;152(2):161–169. 441. Broadley SA, Hartl FU. The role of molecular chaperones in human misfolding diseases. FEBS Lett. 2009;583(16):2647–2653. 442. Ulloa-­Aguirre A, Conn PM. Targeting of G protein-­coupled receptors to the plasma membrane in health and disease. Front Biosci. 2009;14:973–994. 443. Ulloa-­Aguirre A, Janovick JA, Brothers SP, Conn PM. Pharmacologic rescue of conformationally-­defective proteins: implications for the treatment of human disease. Traffic. 2004;5(11):821–837. 444. Hartl FU, Bracher A, Hayer-­Hartl M. Molecular chaperones in protein folding and proteostasis. Nature. 2011;475(7356):324–332. 445. Hutt DM, Powers ET, Balch WE. The proteostasis boundary in misfolding diseases of membrane traffic. FEBS Lett. 2009;583(16):2639–2646. 446. Ron D, Walter P. Signal integration in the endoplasmic Nat Rev Mol Cell Biol.

CHAPTER 2  The Gonadotropin Hormones and Their Receptors 447. Brooks DA. Introduction: molecular chaperones of the ER: their role in protein folding and genetic disease. Semin Cell Dev Biol. 1999;10(5):441–442. 448. Hartl FU, Hayer-­ Hartl M. Molecular chaperones in the cytosol: from nascent chain to folded protein. Science. 2002;295(5561):1852–1858. 449. Morello JP, Petaja-­ Repo UE, Bichet DG, Bouvier M. Pharmacological chaperones: a new twist on receptor folding. Trends Pharmacol Sci. 2000;21(12):466–469. 450. Morello JP, Salahpour A, Laperriere A, et al. Pharmacological chaperones rescue cell-­surface expression and function of misfolded V2 vasopressin receptor mutants. J Clin Invest. 2000;105(7):887–895. 451. Chevet E, Cameron PH, Pelletier MF, Thomas DY, Bergeron JJ. The endoplasmic reticulum: integration of protein folding, quality control, signaling and degradation. Curr Opin Struct Biol. 2001;11(1):120–124. 452. Hou ZS, Ulloa-­Aguirre A, Tao YX. Pharmacoperone drugs: targeting misfolded proteins causing lysosomal storage-­, ion channels-­, and G protein-­coupled receptors-­associated conformational disorders. Expert Rev Clin Pharmacol. 2018;11(6):611–624. 453. Klausner RD, Sitia R. Protein degradation in the endoplasmic reticulum. Cell. 1990;62(4):611–614. 454. Schubert U, Anton LC, Gibbs J, Norbury CC, Yewdell JW, Bennink JR. Rapid degradation of a large fraction of newly synthesized proteins by proteasomes. Nature. 2000;404(6779):770–774. 455. Helenius A, Aebi M. Roles of N-­linked glycans in the endoplasmic reticulum. Annu Rev Biochem. 2004;73:1019–1049. 456. Davis D, Liu X, Segaloff DL. Identification of the sites of N-­linked glycosylation on the follicle-­ stimulating hormone (FSH) receptor and assessment of their role in FSH receptor function. Mol Endocrinol. 1995;9(2):159–170. 457. Clouser CL, Menon KM. N-­linked glycosylation facilitates processing and cell surface expression of rat luteinizing hormone receptor. Mol Cell Endocrinol. 2005;235(1–2):11–19. 458. Mizrachi D, Segaloff DL. Intracellularly located misfolded glycoprotein hormone receptors associate with different chaperone proteins than their cognate wild-­ type receptors. Mol Endocrinol. 2004;18(7):1768–1777. 459. Rozell TG, Davis DP, Chai Y, Segaloff DL. Association of gonadotropin receptor precursors with the protein folding chaperone calnexin. Endocrinology. 1998;139(4):1588–1593. 460. Gething MJ. Role and regulation of the ER chaperone BiP. Semin Cell Dev Biol. 1999;10(5):465–472. 461. Siffroi-­Fernandez S, Giraud A, Lanet J, Franc JL. Association of the thyrotropin receptor with calnexin, calreticulin and BiP. Effects on the maturation of the receptor. Eur J Biochem. 2002;269(20):4930–4937. 462. Zarinan T, Perez-­ Solis MA, Maya-­ Nunez G, et al. Dominant negative effects of human follicle-­ stimulating hormone receptor expression-­ deficient mutants on wild-­ type receptor cell surface expression. Rescue of oligomerization-­ dependent defective receptor expression by using cognate decoys. Mol Cell Endocrinol. 2010;321(2):112–122. 463. Tao YX, Johnson NB, Segaloff DL. Constitutive and agonist-­ dependent self-­association of the cell surface human lutropin receptor. J Biol Chem. 2004;279(7):5904–5914. 464. Nechamen CA, Dias JA. Point mutations in follitropin receptor result in ER retention. Mol Cell Endocrinol. 2003;201(1–2):123–131. 465. Thomas RM, Nechamen CA, Mazurkiewicz JE, Muda M, Palmer S, Dias JA. Follicle-­stimulating hormone receptor forms oligomers and shows evidence of carboxyl-­ terminal proteolytic processing. Endocrinology. 2007;148(5):1987–1995. 466. Lindau-­Shepard B, Brumberg HA, Peterson AJ, Dias JA. Reversible immunoneutralization of human follitropin receptor. J Reprod Immunol. 2001;49(1):1–19. 467. Ulloa-­Aguirre A, Dias JA, Bousfield G, Huhtaniemi I, Reiter E. Trafficking of the follitropin receptor. Meth Enzymol. 2013;521:17–45. 468. Blanpain C, Wittamer V, Vanderwinden JM, et al. Palmitoylation of CCR5 is critical for receptor trafficking and efficient activation of intracellular signaling pathways. J Biol Chem. 2001;276(26):23795–23804. 469. Tanaka K, Nagayama Y, Nishihara E, Namba H, Yamashita S, Niwa M. Palmitoylation of human thyrotropin receptor: slower intracellular trafficking of the palmitoylation-­ defective mutant. Endocrinology. 1998;139(2):803–806.

55.e11

470. Resh MD. Palmitoylation of ligands, receptors, and intracellular signaling molecules. Sci STKE. 2006;2006(359):re14. 471. Fukushima Y, Saitoh T, Anai M, et al. Palmitoylation of the canine histamine H2 receptor occurs at Cys(305) and is important for cell surface targeting. Biochim Biophys Acta. 2001;1539(3):181–191. 472. Percherancier Y, Planchenault T, Valenzuela-­ Fernandez A, Virelizier JL, Arenzana-­Seisdedos F, Bachelerie F. Palmitoylation-­ dependent control of degradation, life span, and membrane expression of the CCR5 receptor. J Biol Chem. 2001;276(34):31936–31944. 473. Kawate N, Menon KM. Palmitoylation of luteinizing hormone/ human choriogonadotropin receptors in transfected cells. Abolition of palmitoylation by mutation of Cys-­621 and Cys-­622 residues in the cytoplasmic tail increases ligand-­induced internalization of the receptor. J Biol Chem. 1994;269(48):30651–30658. 474. Sposini S, Hanyaloglu AC. Spatial encryption of G protein-­coupled receptor signaling in endosomes; Mechanisms and applications. Biochem Pharmacol. 2017;143:1–9. 475. Sposini S, Jean-­ Alphonse FG, Ayoub MA, et al. Integration of GPCR signaling and sorting from very early endosomes via opposing APPL1 mechanisms. Cell Rep. 2017;21(10):2855–2867. 476. Johnson GP, Jonas KC. Mechanistic insight into how gonadotropin hormone receptor complexes direct signalingdagger. Biol Reprod. 2020;102(4):773–783. 477. Hanyaloglu AC, von Zastrow M. Regulation of GPCRs by endocytic membrane trafficking and its potential implications. Annu Rev Pharmacol Toxicol. 2008;48:537–568. 478. Marchese A, Paing MM, Temple BR, Trejo J. G protein-­coupled receptor sorting to endosomes and lysosomes. Annu Rev Pharmacol Toxicol. 2008;48:601–629. 479. Hirakawa T, Galet C, Kishi M, Ascoli M. GIPC binds to the human lutropin receptor (hLHR) through an unusual PDZ domain binding motif, and it regulates the sorting of the internalized human choriogonadotropin and the density of cell surface hLHR. J Biol Chem. 2003;278(49):49348–49357. 480. Kishi M, Liu X, Hirakawa T, Reczek D, Bretscher A, Ascoli M. Identification of two distinct structural motifs that, when added to the C-­terminal tail of the rat LH receptor, redirect the internalized hormone-­receptor complex from a degradation to a recycling pathway. Mol Endocrinol. 2001;15(9):1624–1635. 481. Lin DC, Quevedo C, Brewer NE, et al. APPL1 associates with TrkA and GIPC1 and is required for nerve growth factor-­mediated signal transduction. Mol Cell Biol. 2006;26(23):8928–8941. 482. Irannejad R, Tomshine JC, Tomshine JR, et al. Conformational biosensors reveal GPCR signalling from endosomes. Nature. 2013;495(7442):534–538. 483. Jean-­Alphonse F, Bowersox S, Chen S, Beard G, Puthenveedu MA, Hanyaloglu AC. Spatially restricted G protein-­coupled receptor activity via divergent endocytic compartments. J Biol Chem. 2014;289(7):3960–3977. 484. Marion S, Robert F, Crepieux P, et al. G protein-­coupled receptor kinases and beta arrestins are relocalized and attenuate cyclic 3’,5’-­ adenosine monophosphate response to follicle-­ stimulating hormone in rat primary Sertoli cells. Biol Reprod. 2002;66(1):70–76. 485. Reiter E, Lefkowitz RJ. GRKs and beta-­arrestins: roles in receptor silencing, trafficking and signaling. Trends Endocrinol Metab. 2006;17(4):159–165. 486. Piketty V, Kara E, Guillou F, Reiter E, Crepieux P. Follicle-­ stimulating hormone (FSH) activates extracellular signal-­regulated kinase phosphorylation independently of beta-­ arrestin-­and dynamin-­ mediated FSH receptor internalization. Reprod Biol Endocrinol. 2006;4:33. 487. Mukherjee S, Gurevich VV, Jones JC, et al. The ADP ribosylation factor nucleotide exchange factor ARNO promotes beta-­ arrestin release necessary for luteinizing hormone/choriogonadotropin receptor desensitization. Proc Natl Acad Sci U S A. 2000;97(11):5901–5906. 488. Ahtiainen P, Rulli S, Pakarainen T, Zhang FP, Poutanen M, Huhtaniemi I. Phenotypic characterisation of mice with exaggerated and missing LH/hCG action. Mol Cell Endocrinol. 2007;260–262:255–263. 489. Kumar TR. Gonadotropin gene targeting and biological implications. Endocrine. 2005;26(3):227–233. 490. Pakarainen T, Zhang FP, Nurmi L, Poutanen M, Huhtaniemi I. Knockout of luteinizing hormone receptor abolishes the effects of follicle-­ stimulating hormone on preovulatory maturation and ovulation of mouse Graafian follicles. Mol Endocrinol.

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55.e12 PART I  The Fundamentals of Reproduction 491. Rulli SB, Huhtaniemi I. What have gonadotrophin overexpressing transgenic mice taught us about gonadal function? Reproduction. 2005;130(3):283–291. 492. Abel MH, Wootton AN, Wilkins V, Huhtaniemi I, Knight PG, Charlton HM. The effect of a null mutation in the follicle-­ stimulating hormone receptor gene on mouse reproduction. Endocrinology. 2000;141(5):1795–1803. 493. Dierich A, Sairam MR, Monaco L, et al. Impairing follicle-­ stimulating hormone (FSH) signaling in vivo: targeted disruption of the FSH receptor leads to aberrant gametogenesis and hormonal imbalance. Proc Natl Acad Sci U S A. 1998;95(23):13612–13617. 494. Lei ZM, Mishra S, Zou W, et al. Targeted disruption of luteinizing hormone/human chorionic gonadotropin receptor gene. Mol Endocrinol. 2001;15(1):184–200. 495. Zhang FP, Poutanen M, Wilbertz J, Huhtaniemi I. Normal prenatal but arrested postnatal sexual development of luteinizing hormone receptor knockout (LuRKO) mice. Mol Endocrinol. 2001;15(1):172–183. 496. Chrusciel M, Ponikwicka-­Tyszko D, Wolczynski S, Huhtaniemi I, Rahman NA. Extragonadal FSHR expression and function-­is it real? Front Endocrinol. 2019;10:32. 497. Sun L, Peng Y, Sharrow AC, et al. FSH directly regulates bone mass. Cell. 2006;125(2):247–260. 498. Sun L, Zhang Z, Zhu LL, et al. Further evidence for direct pro-­ resorptive actions of FSH. Biochem Biophys Res Commun. 2010;394(1):6–11. 499. Cannon JG, Kraj B, Sloan G. Follicle-­ stimulating hormone promotes RANK expression on human monocytes. Cytokine. 2011;53(2):141–144. 500. Stilley JA, Guan R, Duffy DM, Segaloff DL. Signaling through FSH receptors on human umbilical vein endothelial cells promotes angiogenesis. J Clin Endocrinol Metab. 2014;99(5):E813–E820. 501. Planeix F, Siraj MA, Bidard FC, et al. Endothelial follicle-­ stimulating hormone receptor expression in invasive breast cancer and vascular remodeling at tumor periphery. J Exp Clin Cancer Res. 2015;34:12. 502. Siraj A, Desestret V, Antoine M, et al. Expression of follicle-­ stimulating hormone receptor by the vascular endothelium in tumor metastases. BMC Cancer. 2013;13:246. 503. Stilley JA, Christensen DE, Dahlem KB, et al. FSH receptor (FSHR) expression in human extragonadal reproductive tissues and the developing placenta, and the impact of its deletion on pregnancy in mice. Biol Reprod. 2014;91(3):74. 504. Ponikwicka-­Tyszko D, Chrusciel M, Stelmaszewska J, et al. Functional expression of FSH receptor in endometriotic lesions. J Clin Endocrinol Metab. 2016;101(7):2905–2914. 505. Song Y, Wang ES, Xing LL, et al. Follicle-­ stimulating hormone induces postmenopausal dyslipidemia through inhibiting hepatic cholesterol metabolism. J Clin Endocrinol Metab. 2016;101(1):254–263. 506. Liu P, Ji Y, Yuen T. Rendina-­Ruedy E, DeMambro VE, Dhawan S, et al. Blocking FSH induces thermogenic adipose tissue and reduces body fat. Nature. 2017;546(7656):107–112. 507. Zaidi M, Lizneva D, Kim SM, et al. FSH, bone mass, body fat, and biological aging. Endocrinology. 2018;159(10):3503–3514. 508. Allan CM, Kalak R, Dunstan CR, et al. Follicle-­stimulating hormone increases bone mass in female mice. Proc Natl Acad Sci U S A. 2010;107(52):22629–22634. 509. Gao J, Tiwari-­Pandey R, Samadfam R, et al. Altered ovarian function affects skeletal homeostasis independent of the action of follicle-­ stimulating hormone. Endocrinology. 2007;148(6):2613–2621. 510. Prior JC. FSH and bone—important physiology or not? Trends Mol Med. 2007;13(1):1–3. 511. Seibel MJ, Dunstan CR, Zhou H, Allan CM, Handelsman DJ. Sex steroids, not FSH, influence bone mass. Cell. 2006;127(6):1079. 512. Williams GR. Hypogonadal bone loss: sex steroids or gonadotropins? Endocrinology. 2007;148(6):2610–2612. 513. Radu A, Pichon C, Camparo P, et al. Expression of follicle-­ stimulating hormone receptor in tumor blood vessels. N Engl J Med. 2010;363(17):1621–1630. 514. Pakarainen T, Ahtiainen P, Zhang FP, Rulli S, Poutanen M, Huhtaniemi I. Extragonadal LH/hCG action—not yet time to rewrite textbooks. Mol Cell Endocrinol. 2007;269(1–2):9–16. 515. Rao CV, Lei ZM. The past, present and future of nongonadal LH/hCG actions in reproductive biology and medicine. Endocrinol. 2007;269(1–2):2–8.

516. Berndt S, Perrier d’Hauterive S, Blacher S, et al. Angiogenic activity of human chorionic gonadotropin through LH receptor activation on endothelial and epithelial cells of the endometrium. FASEB J. 2006;20(14):2630–2632. 517. Cameo P, Szmidt M, Strakova Z, Mavrogianis P, Sharpe-­Timms KL, Fazleabas AT. Decidualization regulates the expression of the endometrial chorionic gonadotropin receptor in the primate. Biol Reprod. 2006;75(5):681–689. 518. Lacroix A, Hamet P, Boutin JM. Leuprolide acetate therapy in luteinizing hormone—dependent Cushing’s syndrome. N Engl J Med. 1999;341(21):1577–1581. 519. Lacroix A, Ndiaye N, Tremblay J, Hamet P. Ectopic and abnormal hormone receptors in adrenal Cushing’s syndrome. Endocr Rev. 2001;22(1):75–110. 520. Bernichtein S, Alevizaki M, Huhtaniemi I. Is the adrenal cortex a target for gonadotropins? Trends Endocrinol Metab. 2008;19(7):231–238. 521. Apaja PM, Aatsinki JT, Rajaniemi HJ, Petaja-­Repo UE. Expression of the mature luteinizing hormone receptor in rodent urogenital and adrenal tissues is developmentally regulated at a posttranslational level. Endocrinology. 2005;146(8):3224–3232. 522. Apaja PM, Harju KT, Aatsinki JT, Petaja-­Repo UE, Rajaniemi HJ. Identification and structural characterization of the neuronal luteinizing hormone receptor associated with sensory systems. J Biol Chem. 2004;279(3):1899–1906. 523. Milligan G. G protein-­coupled receptor dimerisation: molecular basis and relevance to function. Biochim Biophys Acta. 2007;1768(4):825–835. 524. Kleinau G, Muller A, Biebermann H. Oligomerization of GPCRs involved in endocrine regulation. J Mol Endocrinol. 2016;57(1):R59–R80. 525. Horvat RD, Nelson S, Clay CM, Barisas BG, Roess DA. Intrinsically fluorescent luteinizing hormone receptor demonstrates hormone-­driven aggregation. Biochem Biophys Res Commun. 1999;255(2):382–385. 526. Latif R, Graves P, Davies TF. Oligomerization of the human thyrotropin receptor: fluorescent protein-­tagged hTSHR reveals post-­ translational complexes. J Biol Chem. 2001;276(48):45217–45224. 527. Mazurkiewicz JE, Herrick-­ Davis K, Barroso M, et al. Single-­ molecule analyses of fully functional fluorescent protein-­ tagged follitropin receptor reveal homodimerization and specific heterodimerization with lutropin receptor. Biol Reprod. 2015;92(4):100. 528. Jiang X, Fischer D, Chen X, et al. Evidence for follicle-­ stimulating hormone receptor as a functional trimer. J Biol Chem. 2014;289(20):14273–14282. 529. Guan R, Wu X, Feng X, Zhang M, Hebert TE, Segaloff DL. Structural determinants underlying constitutive dimerization of unoccupied human follitropin receptors. Cell Signal. 2010;22(2): 247–256. 530. Sairam MR. Role of carbohydrates in glycoprotein hormone signal transduction. FASEB J. 1989;3(8):1915–1926. 531. Sairam MR, Bhargavi GN. A role for glycosylation of the alpha subunit in transduction of biological signal in glycoprotein hormones. Science. 1985;229(4708):65–67. 532. Meher BR, Dixit A, Bousfield GR, Lushington GH. Glycosylation effects on FSH-­FSHR interaction dynamics: a case study of different FSH glycoforms by molecular dynamics simulations. PLoS One. 2015;10(9):e0137897. 533. Guan R, Feng X, Wu X, et al. Bioluminescence resonance energy transfer studies reveal constitutive dimerization of the human lutropin receptor and a lack of correlation between receptor activation and the propensity for dimerization. J Biol Chem. 2009;284(12):7483–7494. 534. Zhang M, Feng X, Guan R, Hebert TE, Segaloff DL. A cell surface inactive mutant of the human lutropin receptor (hLHR) attenuates signaling of wild-­type or constitutively active receptors via heterodimerization. Cell Signal. 2009;21(11):1663–1671. 535. Ji I, Lee C, Jeoung M, Koo Y, Sievert GA, Ji TH. Trans-­ activation of mutant follicle-­stimulating hormone receptors selectively generates only one of two hormone signals. Mol Endocrinol. 2004;18(4):968–978. 536. Urizar E, Montanelli L, Loy T, et al. Glycoprotein hormone receptors: link between receptor homodimerization and negative cooperativity. EMBO J. 2005;24(11):1954–1964. 537. Jonas KC, Chen S, Virta M, et al. Temporal reprogramming of calcium signalling via crosstalk of gonadotrophin receptors that associate as functionally asymmetric heteromers. Sci Rep.

CHAPTER 2  The Gonadotropin Hormones and Their Receptors 538. Ji I, Lee C, Song Y, Conn PM, Ji TH. Cis-­and trans-­activation of hormone receptors: the LH receptor. Mol Endocrinol. 2002;16(6):1299–1308. 539. Rivero-­Muller A, Chou YY, Ji I, et al. Rescue of defective G protein-­ coupled receptor function in vivo by intermolecular cooperation. Proc Natl Acad Sci U S A. 2010;107(5):2319–2324. 540. Lee C, Ji I, Ryu K, Song Y, Conn PM, Ji TH. Two defective heterozygous luteinizing hormone receptors can rescue hormone action. J Biol Chem. 2002;277(18):15795–15800. 541. Feng X, Zhang M, Guan R, Segaloff DL. Heterodimerization between the lutropin and follitropin receptors is associated with an attenuation of hormone-­ dependent signaling. Endocrinology. 2013;154(10):3925–3930. 542. Casarini L, Lazzaretti C, Paradiso E, et al. Membrane estrogen receptor (GPER) and follicle-­stimulating hormone receptor (FSHR) heteromeric complexes promote human ovarian follicle survival. iScience. 2020;23(12):101812. 543. Pasapera AM, Jimenez-­ Aguilera Mdel P, Chauchereau A, et al. Effects of FSH and 17beta-­ estradiol on the transactivation of estrogen-­regulated promoters and cell proliferation in L cells. J Steroid Biochem Mol Biol. 2005;94(4):289–302. 544. Ulloa-­Aguirre A, Reiter E, Crépieux P. FSH receptor signaling: complexity of interactions and signal diversity. Endocrinology. 2018;159(8):3020–3035. 545. Ulloa-­Aguirre A, Uribe A, Zarinan T, Bustos-­Jaimes I, Perez-­Solis MA, Dias JA. Role of the intracellular domains of the human FSH receptor in G(alphaS) protein coupling and receptor expression. Mol Cell Endocrinol. 2007;260–262:153–162. 546. Arey BJ, Stevis PE, Deecher DC, et al. Induction of promiscuous G protein coupling of the follicle-­stimulating hormone (FSH) receptor: a novel mechanism for transducing pleiotropic actions of FSH isoforms. Mol Endocrinol. 1997;11(5):517–526. 547. Quintana J, Hipkin RW, Sanchez-­Yague J, Ascoli M. Follitropin (FSH) and a phorbol ester stimulate the phosphorylation of the FSH receptor in intact cells. J Biol Chem. 1994;269(12):8772–8779. 548. Zeleznik AJ, Saxena D, Little-­Ihrig L. Protein kinase B is obligatory for follicle-­stimulating hormone-­induced granulosa cell differentiation. Endocrinology. 2003;144(9):3985–3994. 549. Lin YF, Tseng MJ, Hsu HL, Wu YW, Lee YH, Tsai YH. A novel follicle-­ stimulating hormone-­ induced G alpha h/phospholipase C-­delta1 signaling pathway mediating rat sertoli cell Ca2+−influx. Mol Endocrinol. 2006;20(10):2514–2527. 550. Wayne CM, Fan HY, Cheng X, Richards JS. Follicle-­stimulating hormone induces multiple signaling cascades: evidence that activation of Rous sarcoma oncogene, RAS, and the epidermal growth factor receptor are critical for granulosa cell differentiation. Mol Endocrinol. 2007;21(8):1940–1957. 551. Marion S, Kara E, Crepieux P, et al. G protein-­coupled receptor kinase 2 and beta-­ arrestins are recruited to FSH receptor in stimulated rat primary Sertoli cells. J Endocrinol. 2006;190(2):341–350. 552. Nechamen CA, Thomas RM, Dias JA. APPL1, APPL2, Akt2 and FOXO1a interact with FSHR in a potential signaling complex. Mol Cell Endocrinol. 2007;260–262:93–99. 553. Thomas RM, Nechamen CA, Mazurkiewicz JE, Ulloa-­Aguirre A, Dias JA. The adapter protein APPL1 links FSH receptor to inositol 1,4,5-­trisphosphate production and is implicated in intracellular Ca(2+) mobilization. Endocrinology. 2011;152(4):1691–1701. 554. Gonzalez-­Robayna IJ, Falender AE, Ochsner S, Firestone GL, Richards JS. Follicle-­stimulating hormone (FSH) stimulates phosphorylation and activation of protein kinase B (PKB/Akt) and serum and glucocorticoid-­induced kinase (Sgk): evidence for A kinase-­ independent signaling by FSH in granulosa cells. Mol Endocrinol. 2000;14(8):1283–1300. 555. Richards JS, Pangas SA. New insights into ovarian function. Handb Exp Pharmacol. 2010;(198):3–27. 556. Ulloa-­Aguirre A, Crepieux P, Poupon A, Maurel MC, Reiter E. Novel pathways in gonadotropin receptor signaling and biased agonism. Rev Endocr Metab Disord. 2011;12(4):259–274. 557. Donadeu FX, Ascoli M. The differential effects of the gonadotropin receptors on aromatase expression in primary cultures of immature rat granulosa cells are highly dependent on the density of receptors expressed and the activation of the inositol phosphate cascade. Endocrinology. 2005;146(9):3907–3916.

55.e13

558. Musnier A, Heitzler D, Boulo T, et al. Developmental regulation of p70 S6 kinase by a G protein-­coupled receptor dynamically modelized in primary cells. Cell Mol Life Sci. 2009;66(21):3487–3503. 559. Herndon MK, Law NC, Donaubauer EM, Kyriss B, Hunzicker-­ Dunn M. Forkhead box O member FOXO1 regulates the majority of follicle-­stimulating hormone responsive genes in ovarian granulosa cells. Mol Cell Endocrinol. 2016;434:116–126. 560. Hunzicker-­Dunn ME, Lopez-­Biladeau B, Law NC, Fiedler SE, Carr DW, Maizels ET. PKA and GAB2 play central roles in the FSH signaling pathway to PI3K and AKT in ovarian granulosa cells. Proc Natl Acad Sci U S A. 2012;109(44):E2979–E2988. 561. Law NC, White MF, Hunzicker-­Dunn ME. G protein-­coupled receptors (GPCRs) that signal via protein kinase A (PKA) cross-­ talk at insulin receptor substrate 1 (IRS1) to activate the phosphatidylinositol 3-­ kinase (PI3K)/AKT pathway. J Biol Chem. 2016;291(53):27160–27169. 562. Choi JH, Chen CL, Poon SL, Wang HS, Leung PC. Gonadotropin-­ stimulated epidermal growth factor receptor expression in human ovarian surface epithelial cells: involvement of cyclic AMP-­ dependent exchange protein activated by cAMP pathway. Endocr Relat Cancer. 2009;16(1):179–188. 563. Nechamen CA, Thomas RM, Cohen BD, et al. Human follicle-­ stimulating hormone (FSH) receptor interacts with the adaptor protein APPL1 in HEK 293 cells: potential involvement of the PI3K pathway in FSH signaling. Biol Reprod. 2004;71(2):629–636. 564. Cunningham MA, Zhu Q, Unterman TG, Hammond JM. Follicle-­ stimulating hormone promotes nuclear exclusion of the forkhead transcription factor FoxO1a via phosphatidylinositol 3-­ kinase in porcine granulosa cells. Endocrinology. 2003;144(12):5585–5594. 565. Donaubauer EM, Hunzicker-­ Dunn ME. Extracellular signal-­ regulated kinase (ERK)-­ dependent phosphorylation of Y-­ box-­ binding protein 1 (YB-­1) enhances gene expression in granulosa cells in response to follicle-­stimulating hormone (FSH). J Biol Chem. 2016;291(23):12145–12160. 566. Donaubauer EM, Law NC, Hunzicker-­ Dunn ME. Follicle-­ stimulating hormone (FSH) dependent regulation of extracellular regulated kinase (ERK) phosphorylation by MAP kinase phosphatase MKP3. J Biol Chem. 2016;291(37):19701-19712. 567. Crepieux P, Marion S, Martinat N, et al. The ERK-­dependent signalling is stage-­specifically modulated by FSH, during primary Sertoli cell maturation. Oncogene. 2001;20(34):4696–4709. 568. Cottom J, Salvador LM, Maizels ET, et al. Follicle-­stimulating hormone activates extracellular signal-­ regulated kinase but not extracellular signal-­ regulated kinase kinase through a 100-­ kDa phosphotyrosine phosphatase. J Biol Chem. 2003;278(9):7167–7179. 569. Levy FO, Gudermann T, Perez-­Reyes E, Birnbaumer M, Kaumann AJ, Birnbaumer L. Molecular cloning of a human serotonin receptor (S12) with a pharmacological profile resembling that of the 5-­HT1D subtype. J Biol Chem. 1992;267(11):7553–7562. 570. Gudermann T, Birnbaumer M, Birnbaumer L. Evidence for dual coupling of the murine luteinizing hormone receptor to adenylyl cyclase and phosphoinositide breakdown and Ca2+ mobilization. Studies with the cloned murine luteinizing hormone receptor expressed in L cells. J Biol Chem. 1992;267(7):4479–4488. 571. Andric N, Ascoli M. The luteinizing hormone receptor-­activated extracellularly regulated kinase-­ 1/2 cascade stimulates epiregulin release from granulosa cells. Endocrinology. 2008;149(11):5549–5556. 572. Conti M, Hsieh M, Zamah AM, Oh JS. Novel signaling mechanisms in the ovary during oocyte maturation and ovulation. Mol Cell Endocrinol. 2012;356(1–2):65–73. 573. Fan HY, Liu Z, Mullany LK, Richards JS. Consequences of RAS and MAPK activation in the ovary: the good, the bad and the ugly. Mol Cell Endocrinol. 2012;356(1–2):74–79. 574. Fan HY, Liu Z, Shimada M, et al. MAPK3/1 (ERK1/2) in ovarian granulosa cells are essential for female fertility. Science. 2009;324(5929):938–941. 575. Hernandez-­Gonzalez I, Gonzalez-­Robayna I, Shimada M, et al. Gene expression profiles of cumulus cell oocyte complexes during ovulation reveal cumulus cells express neuronal and immune-­related genes: does this expand their role in the ovulation process? Mol Endocrinol. 2006;20(6):1300–1321. 576. Hsieh M, Lee D, Panigone S, et al. Luteinizing hormone-­dependent activation of the epidermal growth factor network is essential for ovulation. Mol Cell Biol. 2007;27(5):1914–1924.

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55.e14 PART I  The Fundamentals of Reproduction 577. Nautiyal J, Steel JH, Rosell MM, et al. The nuclear receptor cofactor receptor-­interacting protein 140 is a positive regulator of amphiregulin expression and cumulus cell-­oocyte complex expansion in the mouse ovary. Endocrinology. 2010;151(6):2923–2932. 578. Hsieh M, Thao K, Conti M. Genetic dissection of epidermal growth factor receptor signaling during luteinizing hormone-­ induced oocyte maturation. PLoS One. 2011;6(6):e21574. 579. Norris RP, Freudzon M, Mehlmann LM, et al. Luteinizing hormone causes MAP kinase-­dependent phosphorylation and closure of connexin 43 gap junctions in mouse ovarian follicles: one of two paths to meiotic resumption. Development. 2008;135(19):3229–3238. 580. Norris RP, Freudzon M, Nikolaev VO, Jaffe LA. Epidermal growth factor receptor kinase activity is required for gap junction closure and for part of the decrease in ovarian follicle cGMP in response to LH. Reproduction. 2010;140(5):655–662. 581. Norris RP, Ratzan WJ, Freudzon M, et al. Cyclic GMP from the surrounding somatic cells regulates cyclic AMP and meiosis in the mouse oocyte. Development. 2009;136(11):1869–1878. 582. Vaccari S, Weeks 2nd JL, Hsieh M, Menniti FS, Conti M. Cyclic GMP signaling is involved in the luteinizing hormone-­ dependent meiotic maturation of mouse oocytes. Biol Reprod. 2009;81(3):595–604. 583. Matzkin ME, Yamashita S, Ascoli M. The ERK1/2 pathway regulates testosterone synthesis by coordinately regulating the expression of steroidogenic genes in Leydig cells. Mol Cell Endocrinol. 2013;370(1–2):130–137. 584. Yamashita S, Tai P, Charron J, Ko C, Ascoli M. The Leydig cell MEK/ERK pathway is critical for maintaining a functional population of adult Leydig cells and for fertility. Mol Endocrinol. 2011;25(7):1211–1222. 585. Fan HY, Liu Z, Johnson PF, Richards JS. CCAAT/enhancer-­ binding proteins (C/EBP)-­alpha and -­beta are essential for ovulation, luteinization, and the expression of key target genes. Mol Endocrinol. 2011;25(2):253–268. 586. Fan HY, Richards JS. Minireview: physiological and pathological actions of RAS in the ovary. Mol Endocrinol. 2010;24(2):286–298. 587. Rehnitz J, Alcoba DD, Brum IS, et al. FMR1 and AKT/mTOR signalling pathways: potential functional interactions controlling folliculogenesis in human granulosa cells. Reprod Biomed Online. 2017;35(5):485–493. 588. Alam H, Maizels ET, Park Y, et al. Follicle-­stimulating hormone activation of hypoxia-­inducible factor-­1 by the phosphatidylinositol 3-­kinase/AKT/Ras homolog enriched in brain (Rheb)/mammalian target of rapamycin (mTOR) pathway is necessary for induction of select protein markers of follicular differentiation. J Biol Chem. 2004;279(19):19431–19440. 589. Kayampilly PP, Menon KM. Follicle-­stimulating hormone increases tuberin phosphorylation and mammalian target of rapamycin signaling through an extracellular signal-­regulated kinase-­dependent pathway in rat granulosa cells. Endocrinology. 2007;148(8):3950–3957. 590. Papaioannou MD, Pitetti JL, Ro S, et al. Sertoli cell Dicer is essential for spermatogenesis in mice. Dev Biol. 2009;326(1):250–259. 591. Nicholls PK, Harrison CA, Walton KL, McLachlan RI, O’Donnell L, Stanton PG. Hormonal regulation of sertoli cell micro-­RNAs at spermiation. Endocrinology. 2011;152(4):1670–1683. 592. Leon K, Gallay N, Poupon A, Reiter E, Dalbies-­Tran R, Crepieux P. Integrating microRNAs into the complexity of gonadotropin signaling networks. Front Cell Dev Biol. 2013;1:3. 593. Jiajie T, Yanzhou Y, Hoi-­ Hung AC, Zi-­ Jiang C, Wai-­ Yee C. Conserved miR-­10 family represses proliferation and induces apoptosis in ovarian granulosa cells. Sci Rep. 2017;7:41304. 594. Yao N, Yang BQ, Liu Y, et al. Follicle-­stimulating hormone regulation of microRNA expression on progesterone production in cultured rat granulosa cells. Endocrine. 2010;38(2):158–166. 595. Zhang L, Zhang X, Zhang X, Lu Y, Li L, Cui S. MiRNA-­143 mediates the proliferative signaling pathway of FSH and regulates estradiol production. J Endocrinol. 2017;234(1):1–14. 596. Davis JS, Weakland LL, West LA, Farese RV. Luteinizing hormone stimulates the formation of inositol trisphosphate and cyclic AMP in rat granulosa cells. Evidence for phospholipase C generated second messengers in the action of luteinizing hormone. Biochem J. 1986;238(2):597–604. 597. Davis JS, West LA, Weakland LL, Farese RV. Human chorionic gonadotropin activates the inositol 1,4,5-­trisphosphate-­Ca2+ intracellular signalling system in bovine luteal cells. 1986;208(2):287–291.

598. Veldhuis JD. Mechanisms subserving hormone action in the ovary: role of calcium ions as assessed by steady state calcium exchange in cultured swine granulosa cells. Endocrinology. 1987;120(2):445–449. 599. Breen SM, Andric N, Ping T, et al. Ovulation involves the luteinizing hormone-­dependent activation of gq/11 in granulosa cells. Mol Endocrinol. 2013;27(9):1483–1491. 600. Mehlmann LM, Kalinowski RR, Ross LF, Parlow AF, Hewlett EL, Jaffe LA. Meiotic resumption in response to luteinizing hormone is independent of a Gi family G protein or calcium in the mouse oocyte. Dev Biol. 2006;299(2):345–355. 601. Jagarlamudi K, Liu L, Adhikari D, et al. Oocyte-­specific deletion of Pten in mice reveals a stage-­specific function of PTEN/PI3K signaling in oocytes in controlling follicular activation. PLoS One. 2009;4(7):e6186. 602. Zheng W, Gorre N, Shen Y, et al. Maternal phosphatidylinositol 3-­kinase signalling is crucial for embryonic genome activation and preimplantation embryogenesis. EMBO Rep. 2010;11(11):890–895. 603. Zheng W, Nagaraju G, Liu Z, Liu K. Functional roles of the phosphatidylinositol 3-­ kinases (PI3Ks) signaling in the mammalian ovary. Mol Cell Endocrinol. 2012;356(1–2):24–30. 604. Palaniappan M, Menon KM. Human chorionic gonadotropin stimulates theca-­interstitial cell proliferation and cell cycle regulatory proteins by a cAMP-­dependent activation of AKT/mTORC1 signaling pathway. Mol Endocrinol. 2010;24(9):1782–1793. 605. Palaniappan M, Menon KM. Luteinizing hormone/human chorionic gonadotropin-­ mediated activation of mTORC1 signaling is required for androgen synthesis by theca-­interstitial cells. Mol Endocrinol. 2012;26(10):1732–1742. 606. Casarini L, Santi D, Brigante G, Simoni M. Two hormones for one receptor: evolution, biochemistry, actions, and pathophysiology of LH and hCG. Endocr Rev. 2018;39(5):549–592. 607. Riccetti L, Yvinec R, Klett D, et al. Human luteinizing hormone and chorionic gonadotropin display biased agonism at the LH and LH/CG receptors. Sci Rep. 2017;7(1):940. 608. Menon KM, Munshi UM, Clouser CL, Nair AK. Regulation of luteinizing hormone/human chorionic gonadotropin receptor expression: a perspective. Biol Reprod. 2004;70(4):861–866. 609. Richards JS, Russell DL, Ochsner S, Espey LL. Ovulation: new dimensions and new regulators of the inflammatory-­like response. Annu Rev Physiol. 2002;64:69–92. 610. Yang LK, Hou ZS, Tao YX. Biased signaling in naturally occurring mutations of G protein-­coupled receptors associated with diverse human diseases. Biochim Biophys Acta, Mol Basis Dis. 2021;1867(1):165973. 611. Luttrell LM, Maudsley S, Bohn LM. Fulfilling the Promise of “biased” G protein-­ coupled receptor agonism. Mol Pharmacol. 2015;88(3):579–588. 612. Reiter E, Ahn S, Shukla AK, Lefkowitz RJ. Molecular mechanism of beta-­arrestin-­biased agonism at seven-­transmembrane receptors. Annu Rev Pharmacol Toxicol. 2012;52:179–197. 613. Shenoy SK, Drake MT, Nelson CD, et al. beta-­arrestin-­dependent, G protein-­independent ERK1/2 activation by the beta2 adrenergic receptor. J Biol Chem. 2006;281(2):1261–1273. 614. Wei H, Ahn S, Shenoy SK, et al. Independent beta-­arrestin 2 and G protein-­mediated pathways for angiotensin II activation of extracellular signal-­regulated kinases 1 and 2. Proc Natl Acad Sci U S A. 2003;100(19):10782–10787. 615. Timossi CM, Barrios de Tomasi J, Zambrano E, Gonzalez R, Ulloa-­ Aguirre A. A naturally occurring basically charged human follicle-­ stimulating hormone (FSH) variant inhibits FSH-­induced androgen aromatization and tissue-­type plasminogen activator enzyme activity in vitro. Neuroendocrinology. 1998;67(3):153–163. 616. Timossi CM, Barrios-­de-­Tomasi J, Gonzalez-­ Suarez R, et al. Differential effects of the charge variants of human follicle-­ stimulating hormone. J Endocrinol. 2000;165(2):193–205. 617. Landomiel F, De Pascali F, Raynaud P, et al. Biased signaling and allosteric modulation at the FSHR. Front Endocrinol (Lausanne). 2019;10:148. 618. Arey BJ, Lopez FJ. Are circulating gonadotropin isoforms naturally occurring biased agonists? Basic and therapeutic implications. Rev Endocr Metab Disord. 2011;12(4):275–288. 619. Dias JA, Bonnet B, Weaver BA, et al. A negative allosteric modulator demonstrates biased antagonism of the follicle stimulating hormone receptor. Mol Cell Endocrinol. 2011;333(2):143–150. 620. Wehbi V, Decourtye J, Piketty V, Durand G, Reiter E, Maurel MC. Selective modulation of follicle-­stimulating hormone signaling pathways with enhancing equine chorionic gonadotropin/antibody immune complexes. Endocrinology. 2010;151(6):2788–2799.

CHAPTER 2  The Gonadotropin Hormones and Their Receptors 621. Wehbi V, Tranchant T, Durand G, et al. Partially deglycosylated equine LH preferentially activates beta-­arrestin-­dependent signaling at the follicle-­stimulating hormone receptor. Mol Endocrinol. 2010;24(3):561–573. 622. Butnev VY, Singh V, Nguyen VT, Bousfield GR. Truncated equine LH beta and asparagine(56)-­ deglycosylated equine LH alpha combine to produce a potent FSH antagonist. J Endocrinol. 2002;172(3):545–555. 623. Aittomaki K, Lucena JL, Pakarinen P, et al. Mutation in the follicle-­ stimulating hormone receptor gene causes hereditary hypergonadotropic ovarian failure. Cell. 1995;82(6):959–968. 624. Tranchant T, Durand G, Gauthier C, et al. Preferential beta-­ arrestin signalling at low receptor density revealed by functional characterization of the human FSH receptor A189 V mutation. Mol Cell Endocrinol. 2011;331(1):109–118. 625. Uchida S, Uchida H, Maruyama T, et al. Molecular analysis of a mutated FSH receptor detected in a patient with spontaneous ovarian hyperstimulation syndrome. PLoS One. 2013;8(9):e75478. 626. Zarinan T, Mayorga J, Jardon-­Valadez E, et al. A novel mutation in the FSH receptor (I423T) affecting receptor activation and leading to primary ovarian failure. J Clin Endocrinol Metab. 2021;106(2):e534–e550. 627. Ayoub MA, Yvinec R, Jegot G, et al. Profiling of FSHR negative allosteric modulators on LH/CGR reveals biased antagonism with implications in steroidogenesis. Mol Cell Endocrinol. 2016;436:10–22. 628. Arnhold IJ, Lofrano-­Porto A, Latronico AC. Inactivating mutations of luteinizing hormone beta-­subunit or luteinizing hormone receptor cause oligo-­amenorrhea and infertility in women. Horm Res. 2009;71(2):75–82. 629. Desai SS, Roy BS, Mahale SD. Mutations and polymorphisms in FSH receptor: functional implications in human reproduction. Reproduction. 2013;146(6):R235–R248. 630. Huhtaniemi I, Alevizaki M. Mutations along the hypothalamic-­ pituitary-­gonadal axis affecting male reproduction. Reprod Biomed Online. 2007;15(6):622–632. 631. Segaloff D. Diseases associated with mutations of the human lutropin receptors. In: Ya-­Xiong Tao, ed. Progress in Molecular Biology and Translational Science. Academic Press, San Diego CA; 2009:97–114. 632. Ulloa-­Aguirre A, Zariñán T, Dias JA, Conn PM. Mutations in G protein-­ coupled receptors that impact receptor trafficking and reproductive function. Mol Cell Endocrinol. 2014;382(1):411–423. 633. Yuan P, He Z, Zheng L, et al. Genetic evidence of “genuine” empty follicle syndrome: a novel effective mutation in the LHCGR gene and review of the literature. Hum Reprod. 2017;32(4):944–953. 634. Chen C, Xu X, Kong L, et al. Novel homozygous nonsense mutations in LHCGR lead to empty follicle syndrome and 46, XY disorder of sex development. Hum Reprod. 2018;33(7):1364–1369. 635. Tapanainen JS, Aittomaki K, Min J, Vaskivuo T, Huhtaniemi IT. Men homozygous for an inactivating mutation of the follicle-­ stimulating hormone (FSH) receptor gene present variable suppression of spermatogenesis and fertility. Nat Genet. 1997;15(2):205–206. 636. Beau I, Touraine P, Meduri G, et al. A novel phenotype related to partial loss of function mutations of the follicle stimulating hormone receptor. J Clin Invest. 1998;102(7):1352–1359. 637. Aittomaki K, Herva R, Stenman UH, et al. Clinical features of primary ovarian failure caused by a point mutation in the follicle-­ stimulating hormone receptor gene. J Clin Endocrinol Metab. 1996;81(10):3722–3726. 638. Latronico AC, Chai Y, Arnhold IJ, Liu X, Mendonca BB, Segaloff DL. A homozygous microdeletion in helix 7 of the luteinizing hormone receptor associated with familial testicular and ovarian resistance is due to both decreased cell surface expression and impaired effector activation by the cell surface receptor. Mol Endocrinol. 1998;12(3):442–450. 639. Martens JW, Verhoef-­Post M, Abelin N, et al. A homozygous mutation in the luteinizing hormone receptor causes partial Leydig cell hypoplasia: correlation between receptor activity and phenotype. Mol Endocrinol. 1998;12(6):775–784. 640. Richter-­Unruh A, Martens JW, Verhoef-­Post M, et al. Leydig cell hypoplasia: cases with new mutations, new polymorphisms and cases without mutations in the luteinizing hormone receptor gene. Clin Endocrinol. 2002;56(1):103–112. 641. Newton CL, Anderson RC, Kreuchwig A, Krause G, Katz AA, Millar RP. Rescue of function of mutant luteinising hormone receptors with deficiencies in cell surface expression, hormone binding and hormone signalling. Neuroendocrinology

55.e15

642. Newton CL, Anderson RC, Katz AA, Millar RP. Loss-­of-­ Function mutations in the human luteinizing hormone receptor predominantly cause intracellular retention. Endocrinology. 2016;157(11):4364–4377. 643. Lu X, Yan Z, Cai R, et al. Pregnancy and live birth in women with pathogenic LHCGR variants using their own oocytes. J Clin Endocrinol Metab. 2019;104(12):5877–5892. 644. Newton CL, Anderson RC. Pharmacoperones for misfolded gonadotropin receptors. Handb Exp Pharmacol. 2018;245:111–134. 645. Charmandari E, Guan R, Zhang M, et al. Misfolding ectodomain mutations of the lutropin receptor increase efficacy of hormone stimulation. Mol Endocrinol. 2016;30(1):62–76. 646. Gromoll J, Schulz A, Borta H, et al. Homozygous mutation within the conserved Ala-­Phe-­Asn-­Glu-­Thr motif of exon 7 of the LH receptor causes male pseudohermaphroditism. Eur J Endocrinol. 2002;147(5):597–608. 647. Martens JW, Lumbroso S, Verhoef-­Post M, et al. Mutant luteinizing hormone receptors in a compound heterozygous patient with complete Leydig cell hypoplasia: abnormal processing causes signaling deficiency. J Clin Endocrinol Metab. 2002;87(6):2506–2513. 648. Richter-­Unruh A, Wessels HT, Menken U, et al. Male LH-­ independent sexual precocity in a 3.5-­year-­old boy caused by a somatic activating mutation of the LH receptor in a Leydig cell tumor. J Clin Endocrinol Metab. 2002;87(3):1052–1056. 649. Toledo SP, Brunner HG, Kraaij R, et al. An inactivating mutation of the luteinizing hormone receptor causes amenorrhea in a 46,XX female. J Clin Endocrinol Metab. 1996;81(11):3850–3854. 650. Vezzoli V, Duminuco P, Vottero A, et al. A new variant in signal peptide of the human luteinizing hormone receptor (LHCGR) affects receptor biogenesis causing Leydig cell hypoplasia. Hum Mol Genet. 2015;24(21):6003–6012. 651. Wu SM, Hallermeier KM, Laue L, et al. Inactivation of the luteinizing hormone/chorionic gonadotropin receptor by an insertional mutation in Leydig cell hypoplasia. Mol Endocrinol. 1998;12(11):1651–1660. 652. Kremer H, Kraaij R, Toledo SP, et al. Male pseudohermaphroditism due to a homozygous missense mutation of the luteinizing hormone receptor gene. Nat Genet. 1995;9(2):160–164. 653. Latronico AC, Anasti J, Arnhold IJ, et al. Brief report: testicular and ovarian resistance to luteinizing hormone caused by inactivating mutations of the luteinizing hormone-­receptor gene. N Engl J Med. 1996;334(8):507–512. 654. Laue LL, Wu SM, Kudo M, et al. Compound heterozygous mutations of the luteinizing hormone receptor gene in Leydig cell hypoplasia. Mol Endocrinol. 1996;10(8):987–997. 655. Newton CL, Whay AM, McArdle CA, et al. Rescue of expression and signaling of human luteinizing hormone G protein-­coupled receptor mutants with an allosterically binding small-­molecule agonist. Proc Natl Acad Sci U S A. 2011;108(17):7172–7176. 656. Rivero-­Muller A, Potorac I, Pintiaux A, et al. A novel inactivating mutation of the LH/chorionic gonadotrophin receptor with impaired membrane trafficking leading to Leydig cell hypoplasia type 1. Eur J Endocrinol. 2015;172(6):K27–K36. 657. Gromoll J, Eiholzer U, Nieschlag E, Simoni M. Male hypogonadism caused by homozygous deletion of exon 10 of the luteinizing hormone (LH) receptor: differential action of human chorionic gonadotropin and LH. J Clin Endocrinol Metab. 2000;85(6):2281–2286. 658. Muller T, Gromoll J, Simoni M. Absence of exon 10 of the human luteinizing hormone (LH) receptor impairs LH, but not human chorionic gonadotropin action. J Clin Endocrinol Metab. 2003;88(5):2242–2249. 659. Muller T, Gromoll J, Simula AP, Norman R, Sandhowe-­Klaverkamp R, Simoni M. The carboxyterminal peptide of chorionic gonadotropin facilitates activation of the marmoset LH receptor. Exp Clin Endocrinol Diabetes. 2004;112(10):574–579. 660. Zhang FP, Rannikko AS, Manna PR, Fraser HM, Huhtaniemi IT. Cloning and functional expression of the luteinizing hormone receptor complementary deoxyribonucleic acid from the marmoset monkey testis: absence of sequences encoding exon 10 in other species. Endocrinology. 1997;138(6):2481–2490. 661. Rosenthal SM, Grumbach MM, Kaplan SL. Gonadotropin-­independent familial sexual precocity with premature Leydig and germinal cell maturation (familial testotoxicosis): effects of a potent luteinizing hormone-­ releasing factor agonist and medroxyprogesterone acetate therapy in four J Clin Endocrinol Metab. 1983;57(3):571–579.

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55.e16 PART I  The Fundamentals of Reproduction 662. Schedewie HK, Reiter EO, Beitins IZ, et al. Testicular Leydig cell hyperplasia as a cause of familial sexual precocity. J Clin Endocrinol Metab. 1981;52(2):271–278. 663. Laue L, Chan WY, Hsueh AJ, et al. Genetic heterogeneity of constitutively activating mutations of the human luteinizing hormone receptor in familial male-­limited precocious puberty. Proc Natl Acad Sci U S A. 1995;92(6):1906–1910. 664. Laue L, Wu SM, Kudo M, et al. Heterogeneity of activating mutations of the human luteinizing hormone receptor in male-­limited precocious puberty. Biochem Mol Med. 1996;58(2):192–198. 665. Gromoll J, Partsch CJ, Simoni M, et al. A mutation in the first transmembrane domain of the lutropin receptor causes male precocious puberty. J Clin Endocrinol Metab. 1998;83(2):476–480. 666. Evans BA, Bowen DJ, Smith PJ, Clayton PE, Gregory JW. A new point mutation in the luteinising hormone receptor gene in familial and sporadic male limited precocious puberty: genotype does not always correlate with phenotype. J Med Genet. 1996;33(2):143–147. 667. Hai L, McGee SR, Rabideau AC, Paquet M, Narayan P. Infertility in female mice with a gain-­of-­function mutation in the luteinizing hormone receptor is due to irregular estrous cyclicity, anovulation, hormonal alterations, and polycystic ovaries. Biol Reprod. 2015;93(1):16. 668. McGee SR, Narayan P. Precocious puberty and Leydig cell hyperplasia in male mice with a gain of function mutation in the LH receptor gene. Endocrinology. 2013;154(10):3900–3913. 669. Shenker A, Laue L, Kosugi S, Merendino Jr JJ, Minegishi T, Cutler Jr GB. A constitutively activating mutation of the luteinizing hormone receptor in familial male precocious puberty. Nature. 1993;365(6447):652–654. 670. Yano K, Hidaka A, Saji M, et al. A sporadic case of male-­limited precocious puberty has the same constitutively activating point mutation in luteinizing hormone/choriogonadotropin receptor gene as familial cases. J Clin Endocrinol Metab. 1994;79(6):1818–1823. 671. Yano K, Kohn LD, Saji M, Kataoka N, Okuno A, Cutler Jr GB. A case of male-­limited precocious puberty caused by a point mutation in the second transmembrane domain of the luteinizing hormone choriogonadotropin receptor gene. Biochem Biophys Res Commun. 1996;220(3):1036–1042. 672. Cocco S, Meloni A, Marini MG, Cao A, Moi P. A missense (T577I) mutation in the luteinizing hormone receptor gene associated with familial male-­limited precocious puberty. Hum Mutat. 1996;7(2):164–166. 673. Kosugi S, Van Dop C, Geffner ME, et al. Characterization of heterogeneous mutations causing constitutive activation of the luteinizing hormone receptor in familial male precocious puberty. Hum Mol Genet. 1995;4(2):183–188. 674. Ignacak M, Hilczer M, Zarzycki J, Trzeciak WH. Substitution of M398T in the second transmembrane helix of the LH receptor in a patient with familial male-­limited precocious puberty. Endocr J. 2000;47(5):595–599. 675. Kraaij R, Post M, Kremer H, et al. A missense mutation in the second transmembrane segment of the luteinizing hormone receptor causes familial male-­limited precocious puberty. J Clin Endocrinol Metab. 1995;80(11):3168–3172. 676. Latronico AC, Abell AN, Arnhold IJ, et al. A unique constitutively activating mutation in third transmembrane helix of luteinizing hormone receptor causes sporadic male gonadotropin-­ independent precocious puberty. J Clin Endocrinol Metab. 1998;83(7):2435–2440. 677. Shinozaki H, Butnev V, Tao YX, Ang KL, Conti M, Segaloff DL. Desensitization of Gs-­coupled receptor signaling by constitutively active mutants of the human lutropin/choriogonadotropin receptor. J Clin Endocrinol Metab. 2003;88(3):1194–1204. 678. Hebrant A, van Staveren WC, Maenhaut C, Dumont JE, Leclere J. Genetic hyperthyroidism: hyperthyroidism due to activating TSHR mutations. Eur J Endocrinol. 2011;164(1):1–9. 679. Kreuchwig A, Kleinau G, Krause G. Research resource: novel structural insights bridge gaps in glycoprotein hormone receptor analyses. Mol Endocrinol. 2013;27(8):1357–1363. 680. Kreuchwig A, Kleinau G, Kreuchwig F, Worth CL, Krause G. Research resource: update and extension of a glycoprotein hormone receptors web application. Mol Endocrinol. 2011;25(4):707–712. 681. Worth CL, Kreuchwig A, Kleinau G, Krause G. GPCR-­SSFE: a comprehensive database of G-­protein-­coupled receptor template predictions and homology models.

682. Latronico AC, Segaloff DL. Insights learned from L457(3.43) R, an activating mutant of the human lutropin receptor. Mol Cell Endocrinol. 2007;260–262:287–293. 683. Ascoli M. Potential Leydig cell mitogenic signals generated by the wild-­ type and constitutively active mutants of the lutropin/choriogonadotropin receptor (LHR). Mol Cell Endocrinol. 2007;260–262:244–248. 684. Hirakawa T, Ascoli M. A constitutively active somatic mutation of the human lutropin receptor found in Leydig cell tumors activates the same families of G proteins as germ line mutations associated with Leydig cell hyperplasia. Endocrinology. 2003;144(9):3872–3878. 685. Hirakawa T, Galet C, Ascoli M. MA-­10 cells transfected with the human lutropin/choriogonadotropin receptor (hLHR): a novel experimental paradigm to study the functional properties of the hLHR. Endocrinology. 2002;143(3):1026–1035. 686. Canto P, Soderlund D, Ramon G, Nishimura E, Mendez JP. Mutational analysis of the luteinizing hormone receptor gene in two individuals with Leydig cell tumors. Am J Med Gen. 2002;108(2):148–152. 687. Liu G, Duranteau L, Carel JC, Monroe J, Doyle DA, Shenker A. Leydig-­ cell tumors caused by an activating mutation of the gene encoding the luteinizing hormone receptor. N Engl J Med. 1999;341(23):1731–1736. 688. Aittomaki K, Tapanainen J, Huhtaniemi I, de la Chapelle A. Inherited primary amenorrhea. The first gynecological disease of Finnish heritage. Duodecim. 1996;112(1):9–11. 689. Bramble MS, Goldstein EH, Lipson A, et al. A novel follicle-­ stimulating hormone receptor mutation causing primary ovarian failure: a fertility application of whole exome sequencing. Hum Reprod. 2016;31(4):905–914. 690. Kotlar TJ, Young RH, Albanese C, Crowley Jr WF, Scully RE, Jameson JL. A mutation in the follicle-­stimulating hormone receptor occurs frequently in human ovarian sex cord tumors. J Clin Endocrinol Metab. 1997;82(4):1020–1026. 691. Meduri G, Touraine P, Beau I, et al. Delayed puberty and primary amenorrhea associated with a novel mutation of the human follicle-­ stimulating hormone receptor: clinical, histological, and molecular studies. J Clin Endocrinol Metab. 2003;88(8):3491–3498. 692. Doherty E, Pakarinen P, Tiitinen A, et al. A Novel mutation in the FSH receptor inhibiting signal transduction and causing primary ovarian failure. J Clin Endocrinol Metab. 2002;87(3): 1151–1155. 693. Nakamura Y, Maekawa R, Yamagata Y, Tamura I, Sugino N. A novel mutation in exon8 of the follicle-­ stimulating hormone receptor in a woman with primary amenorrhea. Gynecol Endocrinol. 2008;24(12):708–712. 694. Achrekar SK, Modi DN, Meherji PK, Patel ZM, Mahale SD. Follicle stimulating hormone receptor gene variants in women with primary and secondary amenorrhea. J Assist Reprod Genet. 2010;27(6):317–326. 695. Allen LA, Achermann JC, Pakarinen P, et al. A novel loss of function mutation in exon 10 of the FSH receptor gene causing hypergonadotrophic hypogonadism: clinical and molecular characteristics. Hum Reprod. 2003;18(2):251–256. 696. Ulloa-­Aguirre A, Zarinan T, Gutierrez-­ Sagal R, Dias JA. Intracellular trafficking of gonadotropin receptors in health and disease. Handb Exp Pharmacol. 2018;245:1–39. 697. Gromoll J, Simoni M, Nordhoff V, Behre HM, De Geyter C, Nieschlag E. Functional and clinical consequences of mutations in the FSH receptor. Mol Cell Endocrinol. 1996;125(1–2):177–182. 698. Liu H, Guo T, Gong Z, et al. Novel FSHR mutations in Han Chinese women with sporadic premature ovarian insufficiency. Mol Cell Endocrinol. 2019;492:110446. 699. Desai SS, Achrekar SK, Sahasrabuddhe KA, et al. Functional characterization of two naturally occurring mutations (Val514Ala and Ala575Val) in follicle-­ stimulating hormone receptor. J Clin Endocrinol Metab. 2015;100(4):E638–E645. 700. Hugon-­Rodin J, Sonigo C, Gompel A, et al. First mutation in the FSHR cytoplasmic tail identified in a non-­pregnant woman with spontaneous ovarian hyperstimulation syndrome. BMC Med Genet. 2017;18(1):44. 701. Rannikko A, Pakarinen P, Manna PR, et al. Functional characterization of the human FSH receptor with an inactivating Ala189Val

CHAPTER 2  The Gonadotropin Hormones and Their Receptors 702. Layman LC, Porto AL, Xie J, et al. FSH beta gene mutations in a female with partial breast development and a male sibling with normal puberty and azoospermia. J Clin Endocrinol Metab. 2002;87(8):3702–3707. 703. Jardon-­Valadez E, Castillo-­Guajardo D, Martinez-­Luis I, Gutierrez-­ Sagal R, Zarinan T, Ulloa-­Aguirre A. Molecular dynamics simulation of the follicle-­stimulating hormone receptor. Understanding the conformational dynamics of receptor variants at positions N680 and D408 from in silico analysis. PLoS One. 2018;13(11):e0207526. 704. Gromoll J, Simoni M, Nieschlag E. An activating mutation of the follicle-­stimulating hormone receptor autonomously sustains spermatogenesis in a hypophysectomized man. J Clin Endocrinol Metab. 1996;81(4):1367–1370. 705. Allan CM, Garcia A, Spaliviero J, Jimenez M, Handelsman DJ. Maintenance of spermatogenesis by the activated human (Asp567Gly) FSH receptor during testicular regression due to hormonal withdrawal. Biol Reprod. 2006;74(5):938–944. 706. Allan CM, Lim P, Robson M, Spaliviero J, Handelsman DJ. Transgenic mutant D567G but not wild-­type human FSH receptor overexpression provides FSH-­ independent and promiscuous glycoprotein hormone Sertoli cell signaling. Am J Physiol Endocrinol Metab. 2009;296(5):E1022–E1028. 707. Haywood M, Tymchenko N, Spaliviero J, et al. An activated human follicle-­stimulating hormone (FSH) receptor stimulates FSH-­like activity in gonadotropin-­deficient transgenic mice. Mol Endocrinol. 2002;16(11):2582–2591. 708. De Leener A, Montanelli L, Van Durme J, et al. Presence and absence of follicle-­stimulating hormone receptor mutations provide some insights into spontaneous ovarian hyperstimulation syndrome physiopathology. J Clin Endocrinol Metab. 2006;91(2):555–562. 709. Montanelli L, Delbaere A, Di Carlo C, et al. A mutation in the follicle-­stimulating hormone receptor as a cause of familial spontaneous ovarian hyperstimulation syndrome. J Clin Endocrinol Metab. 2004;89(3):1255–1258. 710. Smits G, Olatunbosun O, Delbaere A, Pierson R, Vassart G, Costagliola S. Ovarian hyperstimulation syndrome due to a mutation in the follicle-­stimulating hormone receptor. N Engl J Med. 2003;349(8):760–766. 711. Vasseur C, Rodien P, Beau I, et al. A chorionic gonadotropin-­ sensitive mutation in the follicle-­stimulating hormone receptor as a cause of familial gestational spontaneous ovarian hyperstimulation syndrome. N Engl J Med. 2003;349(8):753–759. 712. Nappi RG, Di Naro E, D’Aries AP, Nappi L. Natural pregnancy in hypothyroid woman complicated by spontaneous ovarian hyperstimulation syndrome. Am J Obstet Gynecol. 1998;178(3):610–611. 713. Taher BM, Ghariabeh RA, Jarrah NS, Hadidy AM, Radaideh AM, Ajlouni KM. Spontaneous ovarian hyperstimulation syndrome caused by hypothyroidism in an adult. Eur J Obstet Gynecol Reprod Biol. 2004;112(1):107–109. 714. Kudo M, Osuga Y, Kobilka BK, Hsueh AJ. Transmembrane regions V and VI of the human luteinizing hormone receptor are required for constitutive activation by a mutation in the third intracellular loop. J Biol Chem. 1996;271(37):22470–22478. 715. Zhang M, Tao YX, Ryan GL, Feng X, Fanelli F, Segaloff DL. Intrinsic differences in the response of the human lutropin receptor versus the human follitropin receptor to activating mutations. J Biol Chem. 2007;282(35):25527–25539. 716. Casarini L, Pignatti E, Simoni M. Effects of polymorphisms in gonadotropin and gonadotropin receptor genes on reproductive function. Rev Endocr Metab Disord. 2011;12(4):303–321. 717. Desai SS, Achrekar SK, Paranjape SR, Desai SK, Mangoli VS, Mahale SD. Association of allelic combinations of FSHR gene polymorphisms with ovarian response. Reprod Biomed Online. 2013;27(4):400–406. 718. Brigante G, Spaggiari G, Santi D, et al. The TRHR gene is associated with hypothalamo-­pituitary sensitivity to levothyroxine. Eur Thyroid J. 2014;3(2):101–108. 719. Laan M, Grigorova M, Huhtaniemi IT. Pharmacogenetics of follicle-­stimulating hormone action. Curr Opin Endocrinol Diabetes Obes. 2012;19(3):220–227. 720. Piersma D, Berns EM, Verhoef-­Post M, et al. A common polymorphism renders the luteinizing hormone receptor protein more active by improving signal peptide function and predicts

55.e17

adverse outcome in breast cancer patients. J Clin Endocrinol Metab. 2006;91(4):1470–1476. 721. Piersma D, Verhoef-­Post M, Berns EM, Themmen AP. LH receptor gene mutations and polymorphisms: an overview. Mol Cell Endocrinol. 2007;260–262:282–286. 722. Piersma D, Verhoef-­Post M, Look MP, et al. Polymorphic variations in exon 10 of the luteinizing hormone receptor: functional consequences and associations with breast cancer. Mol Cell Endocrinol. 2007;276(1–2):63–70. 723. Powell BL, Piersma D, Kevenaar ME, et al. Luteinizing hormone signaling and breast cancer: polymorphisms and age of onset. J Clin Endocrinol Metab. 2003;88(4):1653–1657. 724. Simoni M, Tuttelmann F, Michel C, Bockenfeld Y, Nieschlag E, Gromoll J. Polymorphisms of the luteinizing hormone/chorionic gonadotropin receptor gene: association with maldescended testes and male infertility. Pharmacogenet Genom. 2008;18(3): 193–200. 725. García G, Zariñán T, Rodríguez-­Valentín R, et al. Frequency of the 919G>A, 2039A>G, and −29 (G/A) single-­nucleotide polymorphisms (SNPS) in the follicle-­stimulating hormone receptor (FSHR) gene in Mexican Mestizo women. Fertil Steril. 2014;102(3S):e64. 726. Grigorova M, Punab M, Poolamets O, et al. Study in 1790 Baltic men: FSHR Asn680Ser polymorphism affects total testes volume. Andrology. 2013;1(2):293–300. 727. Tranchant T, Durand G, Piketty V, et al. N680S SNP of the human FSH receptor impacts on basal FSH and estradiol level in women and modifies PKA nucelar translocation and CREB-­dependent gene transcription in vitro. Hum Reprod. 2012;27(suppl 1):i45–i46. 728. Hsu SY. New insights into the evolution of the relaxin-­LGR signaling system. Trends Endocrinol Metab. 2003;14(7):303–309. 729. Kong RC, Shilling PJ, Lobb DK, Gooley PR, Bathgate RA. Membrane receptors: structure and function of the relaxin family peptide receptors. Mol Cell Endocrinol. 2010;320(1–2):1–15. 730. Bathgate RA, Halls ML, van der Westhuizen ET, Callander GE, Kocan M, Summers RJ. Relaxin family peptides and their receptors. Physiol Rev. 2013;93(1):405–480. 731. de Lau W, Barker N, Low TY, et al. Lgr5 homologues associate with Wnt receptors and mediate R-­ spondin signalling. Nature. 2011;476(7360):293–297. 732. Feng S, Ferlin A, Truong A, et al. INSL3/RXFP2 signaling in testicular descent. Ann N Y Acad Sci. 2009;1160:197–204. 733. Ferlin A, Pepe A, Gianesello L, et al. New roles for INSL3 in adults. Ann N Y Acad Sci. 2009;1160:215–218. 734. Glinka A, Dolde C, Kirsch N, et al. LGR4 and LGR5 are R-­spondin receptors mediating Wnt/beta-­catenin and Wnt/PCP signalling. EMBO Rep. 2011;12(10):1055–1061. 735. Ivell R, Bathgate RA. Reproductive biology of the relaxin-­like factor (RLF/INSL3). Biol Reprod. 2002;67(3):699–705. 736. Staus DP, Strachan RT, Manglik A, et al. Allosteric nanobodies reveal the dynamic range and diverse mechanisms of G-­protein-­ coupled receptor activation. Nature. 2016;535(7612):448–452. 737. Veber DF, Johnson SR, Cheng HY, Smith BR, Ward KW, Kopple KD. Molecular properties that influence the oral bioavailability of drug candidates. J Med Chem. 2002;45(12):2615–2623. 738. Bonger KM, Hoogendoorn S, van Koppen CJ, Timmers CM, Overkleeft HS, van der Marel GA. Synthesis and pharmacological evaluation of dimeric follicle-­stimulating hormone receptor antagonists. ChemMedChem. 2009;4(12):2098–2102. 739. Arey BJ. Allosteric modulators of glycoprotein hormone receptors: discovery and therapeutic potential. Endocrine. 2008;34:1–10. 740. Arey BJ, Yanofsky SD, Claudia Perez M, et al. Differing pharmacological activities of thiazolidinone analogs at the FSH receptor. Biochem Biophys Res Commun. 2008;368(3):723–728. 741. Guo T. Small molecule agonists and antagonists for the LH and FSH receptors. Expert Opin Ther Pat. 2005;15:1555–1564. 742. Guo T, Adang AE, Dolle RE, et al. Small molecule biaryl FSH receptor agonists. Part 1: lead discovery via encoded combinatorial synthesis. Bioorg Med Chem Lett. 2004;14(7):1713–1716. 743. Guo T, Adang AE, Dong G, et al. Small molecule biaryl FSH receptor agonists. Part 2: lead optimization via parallel synthesis. Bioorg Med Chem Lett. 2004;14(7):1717–1720. 744. Guo T, Adang AE, Dong G, et al. Small molecule biaryl FSH receptor agonists. Part 2: lead optimization via parallel synthesis. Bioorg Med Chem Lett (2004) 14(7):1717–20.

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55.e18 PART I  The Fundamentals of Reproduction 745. Maclean D, Holden F, Davis AM, et al. Agonists of the follicle stimulating hormone receptor from an encoded thiazolidinone library. J Comb Chem. 2004;6(2):196–206. 746. Palmer SS, McKenna S, Arkinstall S. Discovery of new molecules for future treatment of infertility. Reprod Biomed Online. 2005;10:45–54. 747. Nataraja SG, Yu HN, Palmer SS. Discovery and development of small molecule allosteric modulators of glycoprotein hormone receptors. Front Endocrinol. 2015;6:142. 748. Sriraman V, Denis D, de Matos D, Yu H, Palmer S, Nataraja S. Investigation of a thiazolidinone derivative as an allosteric modulator of follicle stimulating hormone receptor: evidence for its ability to support follicular development and ovulation. Biochem Pharmacol. 2014;89(2):266–275. 749. Yu HN, Richardson TE, Nataraja S, et al. Discovery of substituted benzamides as follicle stimulating hormone receptor allosteric modulators. Bioorg Med Chem Lett. 2014;24(9):2168–2172. 750. Heitman LH, Oosterom J, Bonger KM, Timmers CM, Wiegerinck PH, Ijzerman AP. [3H]Org 43553, the first low-­molecular-­weight agonistic and allosteric radioligand for the human luteinizing hormone receptor. Mol Pharmacol. 2008;73(2):518–524. 751.  van de Lagemaat R, Raafs BC, van Koppen C, Timmers CM, Mulders SM, Hanssen RG. Prevention of the onset of ovarian hyperstimulation syndrome (OHSS) in the rat after ovulation induction with a low molecular weight agonist of the LH receptor compared with hCG and rec-­LH. Endocrinology. 2011;152(11):4350–4357. 752.  van de Lagemaat R, Timmers CM, Kelder J, van Koppen C, Mosselman S, Hanssen RG. Induction of ovulation by a potent, orally active, low molecular weight agonist (Org 43553) of the luteinizing hormone receptor. Hum Reprod. 2009;24(3):640–648. 753. Bonger KM, van den Berg RJ, Knijnenburg AD, et al. Discovery of selective luteinizing hormone receptor agonists using the bivalent ligand method. ChemMedChem. 2009;4(7):1189–1195.

754. van Straten NC, van Berkel TH, Demont DR, et al. Identification of substituted 6-­amino-­4-­phenyltetrahydroquinoline derivatives: potent antagonists for the follicle-­stimulating hormone receptor. J Med Chem. 2005;48(6):1697–1700. 755. Arey BJ, Deecher DC, Shen ES, et al. Identification and characterization of a selective, nonpeptide follicle-­stimulating hormone receptor antagonist. Endocrinology. 2002;143(10):3822–3829. 756. Manivannan E, Prasanna S. First QSAR report on FSH receptor antagonistic activity: quantitative investigations on physico-­ chemical and structural features among 6-­ amino-­ 4-­ p henyltetrahydroquinoline derivatives. Bioorg Med Chem Lett. 2005;15(20):4496–4501. 757. Dias JA, Campo B, Weaver BA, et al. Inhibition of follicle-­ stimulating hormone induced preovulatory follicles in rats treated with a nonsteroidal negative allosteric modulator of follicle-­ stimulating hormone receptor. Biol Reprod. 2014;9019:1–11. 758. Janovick JA, Maya-­ Nunez G, Ulloa-­ Aguirre A, et al. Increased plasma membrane expression of human follicle-­stimulating hormone receptor by a small molecule thienopyr(im)idine. Mol Cell Endocrinol. 2009;298(1–2):84–88. 759. Sposini S, De Pascali F, Richardson R, et al. Pharmacological programming of endosomal signaling activated by small molecule ligands of the follicle stimulating hormone receptor. Front Pharmacol. 2020;11:593492. 760. Yanofsky SD, Shen ES, Holden F, et al. Allosteric activation of the follicle-­stimulating hormone (FSH) receptor by selective, nonpeptide agonists. J Biol Chem. 2006;281(19):13226–13233. 761. Duan J, Xu P, Cheng X, et al. Structures of full-­ length glycoprotein hormone receptor signalling complexes. Nature. 2021;598(7882):688–692.

3

Prolactin in Human Reproduction Nicholas A. Tritos

OUTLINE INTRODUCTION LACTOTROPH DEVELOPMENT PROLACTIN GENE PROLACTIN SYNTHESIS IN PITUITARY LACTOTROPHS PROLACTIN SYNTHESIS IN THE DECIDUA AND OTHER TISSUES PROLACTIN ASSAYS Hook Effect Macroprolactin Heterophilic Antibodies Exogenous Biotin PROLACTIN SECRETION Changes in Prolactin Levels With Age Prolactin Levels During Physiologic Stress Prolactin Levels During the Menstrual Cycle, Pregnancy, and the Postpartum State REGULATION OF SYSTEMIC PROLACTIN LEVELS Prolactin Inhibitory Factors Prolactin-­Releasing Factors PROLACTIN ACTIONS Prolactin Receptor Prolactin Effects on the Breast Prolactin Effects on Gonadotropin Secretion Prolactin Effects on the Ovary Prolactin Effects on the Testis Prolactin Effects on the Adrenal Cortex Prolactin and the Skeleton PATHOLOGICAL STATES OF PROLACTIN DEFICIENCY AND EXCESS Prolactin Deficiency Hyperprolactinemia: Causes Hyperprolactinemia: Diagnosis PROLACTIN-­SECRETING PITUITARY ADENOMAS (PROLACTINOMAS) Epidemiology and Natural History Pathogenesis Pathology Clinical Manifestations Management PREGNANCY AND PROLACTINOMAS Prolactin-­Secreting Pituitary Adenomas During Preconception and Pregnancy Effects of Dopamine Agonists During Preconception and Pregnancy Management of Patients With Prolactinomas During Preconception and Pregnancy 56

INTRODUCTION Prolactin (PRL) is a single chain (23 KDa) polypeptide hormone, which is secreted by anterior pituitary lactotroph cells.1 Several lines of evidence indicate that PRL has an essential role in reproduction and lactation.2 In addition, animal data have supported a role for PRL in a variety of metabolic processes.2 However, such PRL actions have not been unequivocally confirmed in humans.3 The present chapter reviews PRL physiology, followed by a discussion of the role of PRL in pathologic states, including hyperprolactinemia and PRL deficiency. Data on the epidemiology, pathology, clinical evaluation, and management of PRL-­secreting pituitary adenomas (prolactinomas) are then reviewed, including data relevant to prolactinomas in the setting of preconception and pregnancy.

LACTOTROPH DEVELOPMENT • L  actotroph cells develop under carefully orchestrated control by several transcription factors. • Lactotroph hyperplasia is physiologic during pregnancy and is reversible postpartum.    Pituitary lactotrophs are relatively abundant in the human anterior pituitary gland, accounting for up to 25% of cells in individuals of both genders.4 During embryogenesis, the pituitary gland develops from ectodermal primordial cells destined to form the anterior and intermediate lobe and neuroectodermal tissue arising from the floor of the diencephalon, which ultimately forms the posterior lobe. During development, inductive interactions and a host of transcription factors have a critical role in the formation of the pituitary and differentiation into mature functioning cells.5–7 Transcription factors and signaling molecules that have been implicated in pituitary ontogenesis include those involved in the initiation of pituitary formation (SIX1, SIX6, HESX1, OTX2, PITX1, PITX2, PITX3, ISL1, LHX3, LHX4, SOX2, beta catenin, NOTCH1, NOTCH2), those involved in the migration and proliferation of cells forming the Rathke’s pouch (BMP2, BMP4, FGF8, FGF10, FGF18, SHH) and those involved in lactotroph differentiation (PROP1, POU1F1, GATA2, LHX3).5–8 Lactotroph and somatotroph cells generally develop from common progenitor cells (mammosomatotrophs) under the influence of several transcription factors, though it is possible that some lactotrophs may develop through other precursor cell lines.9,10 In particular, the transcription factor POU1F1 (also known as PIT1), a member of the POU homeodomain transcription factor family, is critical in the differentiation and proliferation of lactotrophs, somatotrophs and thyrotrophs and the expression of genes encoding PRL, growth hormone (GH) and the beta subunit of thyrotropin (TSH beta).7,9 Patients with inactivating mutations in the POU1F1 gene lack lactotrophs, somatotrophs, and thyrotrophs, resulting in PRL, GH, and thyrotropin deficiency, respectively. In addiPIT1 antibody syndrome develop

CHAPTER 3  Prolactin in Human Reproduction

PRL, GH, and thyrotropin deficiency as a consequence of autoimmune damage to the respective pituitary cell populations.11 Another homeodomain transcription factor, known as prophet of PIT1 (PROP1), has a critical role in the expression of POU1F1.7,9 Patients with inactivating mutations in the PROP1 gene may have PRL, GH, thyrotropin deficiency as well as gonadotropin (follicle stimulating hormone [FSH] and luteinizing hormone [LH]) deficiency. Patients with inactivating mutations in genes encoding other transcription factors, including HESX1, LHX3, and LHX4, generally have multiple pituitary hormone deficiencies as well as other midline cranial defects.7,9

PROLACTIN GENE A •  single gene encodes prolactin in humans. • Prolactin gene transcription is increased by estradiol and inhibited by thyroid hormone. • Dopamine, the major factor regulating prolactin secretion in humans, acts by inhibiting adenyl cyclase–dependent signaling pathways.    In humans, the gene encoding PRL consists of 5 coding exons, one noncoding exon, and four introns. It spans approximately 10 Kb in length and is located on chromosome 6.12,13 There are several regulatory elements located in the 5’ region of the PRL gene, including areas responsible for stimulation of PRL gene transcription in response to POU1F1 or estradiol, as well as those responsible for suppression of PRL gene transcription by thyroid hormone.14–18 In addition to its direct effects, estradiol also modulates dopamine (DA)-­ induced inhibition on PRL gene transcription.19–21 The stimulatory effects of POU1F1 on PRL gene transcription are also subject to modulation by a variety of factors, including cyclic adenosine monophosphate (cAMP), glucocorticoids, estradiol, thyrotropin-­releasing hormone (TRH) and epidermal growth factor (EGF).14–16,22–24 Dopamine is the major inhibitory factor regulating PRL secretion and acts through the D2 dopamine receptor to inhibit adenyl cyclase and the cAMP-­ dependent protein kinase A (PKA) pathway.25 In contrast, TRH stimulates PRL secretion via the phosphoinositide pathway, leading to the activation of membrane calcium channels and the release of calcium ions from the endoplasmic reticulum into the cytoplasm, which in turn activate protein kinase C (PKC), causing the downstream phosphorylation of other proteins, and also bind to calmodulin or topoisomerase II, which have a variety of downstream actions.26 Of note, dopamine inhibits PRL secretion caused by intracellular calcium ion release.27 This dopamine action is antagonized by several calcium channel antagonists in vitro, including verapamil, diltiazem, and nimodipine.28 Interestingly, verapamil causes the opposite effect from what would be predicted by in vitro data; that is, it leads to an increase in PRL levels in humans.29,30 This observation was associated with decreased tuberoinfundibular dopamine release and may involve an effect of verapamil on N-­type calcium channels present in neurons.31 Other types of calcium channel antagonists, including dihydropyridines and benzothiazepines, do not affect PRL levels in vivo.31 Vasoactive intestinal peptide (VIP) stimulates adenyl cyclase, leading to PRL secretion.32 Several factors that stimulate PRL secretion, including TRH, neurotensin, and angiotensin II, may also act via the stimulation of phospholipase A2, leading to the release of arachidonic acid, which causes an increase in calcium influx.33–35 This effect can be antagonized by dopamine and phospholipase A2 inhibitors.33,36

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PROLACTIN SYNTHESIS IN PITUITARY LACTOTROPHS • S  everal posttranslational prolactin modifications occur and influence prolactin bioactivity. • Macroprolactin species are large prolactin aggregates that have decreased bioactivity. • A 16 kDa prolactin fragment has been implicated in the pathogenesis of peripartum cardiomyopathy and preeclampsia.    The PRL gene is transcribed to mRNA, which undergoes processing in the nucleus to yield a mature, 1 Kb mRNA species, encoding a 227 amino acid PRL precursor. This contains a 28 amino acid signal peptide sequence, which is cleaved posttranslationally to yield a 199 amino acid PRL protein.1,13 Additional posttranslational modifications of the PRL molecule include glycosylation, phosphorylation, cleavage, and polymerization. Of note, the large majority (80%–90%) of the circulating PRL is monomeric, whereas about 10% of circulating PRL is dimeric (“big PRL,” molecular mass ∼50 KDa) and approximately 5% of circulating PRL is multimeric (“big big PRL”).37–39 Such high molecular mass PRL species are collectively called “macroprolactin” and may additionally contain bound immunoglobulin.40 Macroprolactin has been reported to exhibit decreased binding to PRL receptors and has lower receptor binding affinity and decreased bioactivity in most, but not all, assays.38,39 Patients with macroprolactinemia, who have elevated total serum PRL as a consequence of elevated multimeric PRL, appear to have normal pituitary-­ gonadal function, likely as a consequence of decreased bioactivity of multimeric PRL species.39,41–43 Cleavage of the 23 KDa PRL species by metalloproteases or cathepsin D may occur in peripheral tissues, leading to the generation of an N-­terminal 16 KDa PRL variant.44 This PRL species has antiangiogenic, proapoptotic, and proinflammatory properties and has been implicated in the pathogenesis of peripartum cardiomyopathy and preeclampsia, based on animal and human data.44–48 Bromocriptine therapy decreases serum PRL and improves cardiac function in women with peripartum cardiomyopathy.49–51

PROLACTIN SYNTHESIS IN THE DECIDUA AND OTHER TISSUES • E  xtrapituitary prolactin secretion occurs in the decidua and other tissues. • Prolactin of decidual origin appears to promote immunological tolerance of the fetus in utero. • The physiologic role of extrapituitary prolactin in other tissues remains incompletely understood in humans.    Several lines of evidence suggest that PRL is synthesized in the decidua.52,53 Very high PRL levels (10–100 times those in maternal serum) have been found in amniotic fluid.52,53 In culture, decidual and chorion cells secrete PRL.54 This PRL species is identical in sequence and activity to pituitary PRL and is expressed under control by an alternative promoter, which is located upstream from the transcription initiation site used in pituitary lactotrophs.55,56 Of note, PRL secretion in the decidua is stimulated by progesterone (either alone or together with estrogen), relaxin, insulin, and insulin-­like growth factor I (IGF-­I) but is not influenced by dopamine agonists or antagonists (in contrast to pituitary PRL).57–60 Decidual PRL appears to have an important role in maintaining pregnancy by downregulating interleukin 6 and 20 alpha hydroxysteroid dehydrogenase.61 Of note, decidual PRL synthesis was reduced in decidual tissue from women who had suffered a miscarriage,

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wherein proinflammatory cytokines were elevated.62 In aggregate, these data suggest that decidual PRL supports pregnancy by promoting immunological tolerance of the fetus in utero.61,62 Extrapituitary PRL expression has been documented in the mammary gland, ovaries, testes, prostate, endothelial cells, brain, skin, adipose tissue, lymphocytes, and cochlea in a variety of animal paradigms.63–67 The physiologic role of extrapituitary PRL synthesis is under investigation.64 It has been proposed that extrapituitary PRL may promote tumorigenesis (including tumor initiation and/or propagation) in the breast and prostate.68–70 This is an area of ongoing debate and uncertainty since some studies have found little PRL expression in human tumors.71 Gain-­of-­function PRL receptor variants do not appear to be associated with a higher risk of breast cancer or fibroadenomas.72 Nevertheless, available data have prompted the development of a variety of inhibitors of PRL receptors or PRL signaling, and their study as potential therapies in human prostate or breast cancer.73–75

PROLACTIN ASSAYS • P  rolactin is commonly measured using two-­site antibody immunoassays. • Several artifacts and conditions may influence the results of these assays, including the hook effect, the presence of macroprolactin, heterophilic antibodies, and biotin.    Serum PRL is currently measured using two-­ site (“sandwich”) immunometric assays, including immunoradiometric (IRMA), chemiluminescent (ICMA), and electrochemiluminescent (ECLIA) platforms, which use a two-­antibody system.76 A “capture” antibody is used to collect the PRL molecules present in the specimen to a solid substrate (such as microparticle beads or coated tubes). Subsequently, a second (“reporter”) antibody attaches to the PRL–capture antibody complex and is used to generate a detection signal through its radiolabeled or chemiluminescent tag.76 Measured PRL levels can vary considerably between different immunoassays.77 Assay interference may occur and should be suspected in the presence of nonlinearity in serially diluted specimens or discrepancy between measured hormone levels and the clinical presentation.76 Several factors and mechanisms, including the “hook effect,” the presence of macroprolactin, heterophilic antibodies, or exogenous biotin, may lead to assay interference.76

Hook Effect The hook effect may occur when serum specimens being assayed contain very high PRL levels, as can be the case in patients with large PRL-­ secreting pituitary macroadenomas.78,79 In these patients, PRL is present in vast stoichiometric excess to the two assay antibodies in the test solution, thus preventing the formation of the heterotrimeric complex (capture antibody– PRL–reporter antibody). The hook effect can cause substantial under-­reporting of PRL levels.78–80 This artifact is clinically significant since patients with adenomas that are thought to be PRL-­ secreting will be generally offered a trial of medical therapy.78 In contrast, failure to diagnose a prolactinoma based on a normal or minimally elevated PRL level in a patient with a large pituitary adenoma may likely lead to a recommendation for surgery rather than medical therapy. In one study, the hook effect was reported in 5.6% of 69 patients with macroadenomas.81 The frequency of hook effect using current assay techniques is uncertain. To detect this possible artifact, PRL measurements should be obtained both in undiluted and diluted (1:100) serum specimens in patients with large macroadenomas.78,79

Macroprolactin Monomeric PRL normally accounts for approximately 90% of circulating PRL, the rest consisting of PRL multimers of varying molecular mass, sometimes associated with immunoglobulin. Multimeric PRL species are collectively termed “macroprolactin” (as already mentioned).37–39 Macroprolactin can be resolved and separated from monomeric PRL by gel filtration.37–39 In routine practice, polyethylene glycol (PEG) precipitation has been used to separate macroprolactin from PRL monomers (only the latter is left in the supernatant after centrifugation).39,82 Of note, hyperprolactinemia can be solely attributed to the presence of macroprolactin only if the monomeric PRL level present in the serum is within the reference range.83 Macroprolactin appears to have decreased bioactivity in some, but not all, bioassays.37,39 Women with macroprolactinemia who were treated with dopamine agonists were reported to experience resolution of galactorrhea (if originally present) but had no consistent improvement in menstrual cyclicity.83,84 Macroprolactinemia should be considered in patients with hyperprolactinemia who lack typical symptoms attributable to PRL excess.43,85 However, the presence of macroprolactinemia does not obviate the need for pituitary imaging, since some patients with pituitary adenomas may exhibit macroprolactinemia.42 Therefore, the clinical utility of assessing macroprolactin in symptomatic patients is limited.

Heterophilic Antibodies Heterophilic antibodies, including human anti-­mouse or other anti-­animal immunoglobulins and the rheumatoid factor autoantibody, may cause assay interference by bridging the capture and reporter antibodies in the absence of analyte (i.e., PRL), thus giving rise to artifactually high PRL levels.86,87 Heterophilic antibodies may also confound many other assays, in addition to PRL immunoassays.87 Using blocking animal sera or heterophilic antibody blocking tubes can help prevent this type of interference.86

Exogenous Biotin Several immunoassay platforms use biotinylated capture antibodies that collect the analyte being assayed onto streptavidin-­ coated microbeads.88 Measurement of hormone levels in patients taking biotin supplements in large doses is subject to possible interference if the immunoassay platform in use employs the biotin-­streptavidin reaction.89 In these cases, ingested biotin may prevent the formation of the streptavidin-­biotin complex in the assay solution, leading to under-­reporting of the analyte being measured by sandwich immunoassays (including PRL) or overreporting of hormones measured using competition assays.89,90 If it is clinically permissible, patients taking biotin supplements should discontinue them for 3 days before serum specimens are obtained for immunoassays.91,92 Alternatively, immunoassay formats that do not use biotinylated antibodies can be used.91,92

PROLACTIN SECRETION P •  rolactin secretion is pulsatile. • Serum prolactin levels rise in many physiologic states, including pregnancy and lactation.    Prolactin secretion is pulsatile, involving approximately 14 secretory peaks per 24 hours, each peak lasting 67 to 76 minutes.93,94 The PRL pulse amplitude increases during slow-­wave sleep, with pulses beginning about 60 minutes after sleep onset.95 In addition, PRL levels increase by 50% to 100% about 30 minutes after meals as a result of stimulation of PRL secretion by

CHAPTER 3  Prolactin in Human Reproduction

amino acids being absorbed postprandially (particularly tyrosine, phenylalanine, and glutamate).96,97

Changes in Prolactin Levels With Age Immediately following delivery, PRL levels in neonates are elevated about 10 times above baseline, presumably as a consequence of high levels of placental estrogen, and decrease toward baseline over several months.98 Subsequently, PRL levels rise somewhat during adolescence.98 In women, PRL levels decrease by approximately 50% within about 2 years after menopause.99 In older men, PRL levels also decline by about 50% in comparison with young adults.100

Prolactin Levels During Physiologic Stress Acutely, exercise leads to an increase in serum PRL levels, which is not sustained long-­term in long-­distance runners.101,102 Other forms of physiologic stress, such as acute illness or injury, also lead to a two-­to-­three-­fold increase in PRL levels, lasting about 1 hour, which is not sustained in patients with prolonged illness.101,103 Women have a more robust prolactin response than men.

Prolactin Levels During the Menstrual Cycle, Pregnancy, and the Postpartum State Prolactin levels are lower in the follicular phase and rise during the luteal phase in some women.104 In addition, PRL and LH secretion are synchronous in the luteal phase, possibly in response to gonadotropin-­releasing hormone (GnRH) stimulation of PRL secretion during that period.105 During pregnancy, PRL levels increase continuously and may reach more than tenfold or higher levels in nonpregnant women.53 This is a consequence of estrogen secretion by the placenta, which leads to lactotroph hyperplasia and stimulates PRL secretion, preparing the mammary gland for lactation postpartum.106,107 Lactotroph hyperplasia is physiologic during pregnancy and is reversible within several months after delivery but is delayed by nursing.106,107 After delivery, basal PRL levels remain elevated in women who are nursing.53,108 In addition, PRL secretory peaks occur rapidly at the time of each suckling event as a result of neurogenic stimulation of PRL secretion.53,108 Within several months, there is a gradual decline in basal PRL levels toward normal as well as a decrease in the amplitude of secretory spikes in response to suckling.53,108 These events occur as a consequence of a gradual decrease in the intensity of breastfeeding, while formula is being introduced into the infant’s diet.53,108 Menses resume as postpartum hyperprolactinemia abates. Nipple stimulation, either acute or chronic (by nipple rings), may cause hyperprolactinemia in some healthy women who are not nursing.108,109

REGULATION OF SYSTEMIC PROLACTIN LEVELS • P  rolactin secretion is primarily under inhibitory control by hypothalamic dopamine. • Thyrotropin-­releasing hormone, vasoactive intestinal peptide, serotonin, and other factors may have some role in stimulating prolactin secretion.    Hypothalamic regulation of PRL secretion is primarily mediated via inhibitory factors (predominantly dopamine), as evidenced by an increase in PRL secretion and systemic PRL levels in patients who suffered pituitary stalk damage.110,111 In addition, several releasing factors may have a role in modulating PRL secretion (Fig. 3.1).

59

Stress

3

Dopamine (–)

Serotonin (+)

Drugs Tranquilizers Antidepressants Antihypertensives

Hypothalamus (?) PRFs VIP TRH PRLrp

Pituitary

PIFs Dopamine GAP (?) Estrogen Pregnancy

Prolactin

Spinal afferent

Breast (suckling stimulus)

Fig. 3.1 Factors and pathways involved in the regulation of prolactin secretion.  GAP, Gonadotropin-­releasing hormone-­ associated peptide (precursor to gonadotropin-­releasing hormone); PIFs, prolactin inhibitory factors; PRFs: prolactin-­releasing factors; PRLrp: prolactin-­releasing peptide; TRH: thyrotropin-­releasing hormone; VIP: vasoactive intestinal peptide (Reproduced from Molitch ME. Disorders of prolactin secretion. Endocrinol Metab Clin North Am. 2001;30:585–610, with permission.)

Prolactin Inhibitory Factors Several lines of evidence indicate that hypothalamic dopamine is the most important PRL inhibitory factor under physiologic conditions.112 Dopamine is present in hypothalamic portal vessels at levels that are sufficient to suppress PRL secretion.113,114 Stimuli that elicit PRL secretion also lead to a decrease in dopamine levels in the hypophyseal portal circulation.115,116 Mice with targeted disruption of D2 receptors, whose absence prevents dopamine action in pituitary lactotrophs, develop hyperprolactinemia, lactotroph hyperplasia, and multiple prolactinomas.117–119 Infusion of dopamine in low doses leads to suppression of PRL secretion in humans.120,121 Estradiol partially reverses the effects of dopamine infusion on PRL secretion.122 Pharmacologic agents that inhibit dopamine receptors lead to hyperprolactinemia in humans.123 Dopamine is synthesized in neurons whose perikarya are located in the dorsal arcuate nucleus and the ventromedial nucleus of the hypothalamus.112 This neuronal pathway is termed the tuberoinfundibular dopamine pathway. Axon terminals originating from these neurons terminate in the median eminence, where dopamine is released, enters the hypophyseal portal system, and traverses the pituitary stalk to reach pituitary lactotrophs in the pars distalis, where dopamine activates D2 receptors to suppress PRL secretion.112 Mice with disrupted PRL genes do not synthesize any PRL and have substantially decreased dopamine in the tuberoinfundibular dopamine pathway, consistent with the existence of a positive effect of secreted PRL on hypothalamic dopamine release, generating a short loop feedback mechanism of secreted PRL that negatively regulates its own secretion.124

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Whether other physiologically relevant PRL inhibitory factors exist is a matter of controversy. Some data suggest that gonadotropin-­ releasing hormone (GnRH)-­ associated protein (GAP), neuromedin U as well as gamma-­ aminobutyric acid (GABA) can suppress PRL in experimental paradigms, but their relevance in humans is unclear.125–128 Animal data have also suggested an autocrine and paracrine role for PRL in regulating its own secretion.129

Prolactin-­Releasing Factors Several factors have been proposed as being stimulatory of PRL secretion (as detailed below). However, their physiologic relevance is uncertain in many cases. Overall, their role appears relatively minor in comparison with the inhibitory role of dopamine on PRL secretion.

Thyrotropin-­Releasing Hormone Thyrotropin-­releasing hormone (TRH) induces PRL secretion in vitro and in vivo in humans when administered in doses that also cause TSH secretion.130,131 On the other hand, targeted disruption of the TRH gene in mice leads to central hypothyroidism but does not influence PRL levels.132 In addition, nipple stimulation during suckling leads to a PRL secretory spike but does not influence TSH levels.133 Thus, it appears that TRH does not have a major role as a releasing factor under normal conditions. It should be noted, however, that patients with primary hypothyroidism have elevated TSH and PRL levels, which are normalized by levothyroxine replacement, suggesting that endogenous TRH may be relevant in PRL regulation in these patients but does not exclude the possibility that changes in dopaminergic tone in the hypothyroid state may also contribute to hyperprolactinemia.134–136 Patients with hyperthyroidism have normal PRL levels but do exhibit an abnormally low PRL response to TRH administration, which is restored after euthyroidism is achieved.134–136

Vasoactive Intestinal Peptide and Peptide Histidine Methionine Vasoactive intestinal peptide (VIP) and peptide histidine methionine are derived from posttranslational processing of the same precursor.137,138 Of note, VIP-­ producing neurons have been localized to the paraventricular hypothalamic nucleus.137 In addition, VIP is expressed in the anterior pituitary.139 Administration of VIP induces PRL synthesis and secretion in vitro and in vivo in humans.140,141 In addition, VIP immunoneutralization or antagonism blocks PRL secretion in various paradigms.142,143 Administration of peptide histidine methionine has led to inconsistent results with regard to PRL secretion in humans.141,144 Overall, the physiologic role of VIP and peptide histidine methionine in the regulation of PRL secretion remains incompletely understood.

Serotonin In experimental animals, administration of either serotonin or 5-­hydroxytryptophan (serotonin precursor) induces PRL secretion.145 Inhibition of serotonin synthesis blunts PRL secretion in response to suckling.146 Similarly, intravenous administration of 5-­hydroxytryptophan in humans triggers PRL secretion.147 Cyproheptadine, a serotonin receptor antagonist, inhibits both the nocturnal PRL surge and the fenfluramine-­induced increase in PRL levels.148,149 Administration of fluoxetine, a serotonin reuptake inhibitor, leads to a mild increase in PRL levels over baseline.150 In aggregate, these data suggest a possible role for

serotonin in the regulation of night-­time PRL secretion as well as PRL release after nipple stimulation.

Opioids In experimental animals, administration of opioid peptides results in an increase in PRL secretion, likely mediated via activation of the mu opioid receptor.151,152 It appears that the effect of opioids on PRL secretion is indirect and is mediated via a decrease in dopamine release in the tuberoinfundibular dopamine pathway. 153

In humans, opioid administration results in PRL secretion, both acutely and chronically.154,155 Of note, administration of naloxone, a mu opioid receptor antagonist, causes an increase in PRL levels during the late follicular and midluteal phase of the menstrual cycle in humans but does not blunt PRL secretion in response to physiologic stress.156,157 Overall, available data suggest that endogenous opioids do not have a major physiologic role in the regulation of PRL secretion.

Other Neuropeptides and Neurotransmitters Growth hormone-­ releasing hormone (GHRH) administration triggers PRL secretion in healthy individuals and repeated GHRH administration leads to increased PRL levels in children with GH neurosecretory dysfunction.158,159 However, it is unclear whether GHRH is physiologically relevant in the regulation of PRL secretion. Gonadotropin-­ releasing hormone (GnRH) induces PRL secretion in vitro and in vivo in humans, including women with subfertility who are undergoing ovulation induction with gonadotropins.160,161 In addition, some healthy women secrete PRL in response to GnRH administration, with the highest proportion of responses noted in women around ovulation.162 In contrast, GnRH administration does not induce PRL secretion in healthy men.163,164 Of note, GnRH leads to PRL secretion in transgender men on estrogen therapy, suggesting that the estrogenic milieu may be essential for GnRH to induce PRL release.163,164 Oxytocin administration triggers PRL secretion in vitro and in vivo in some experimental paradigms.165,166 In contrast, oxytocin alone does not appear to affect PRL secretion in humans.167 Oxytocin administration has a minor positive effect on VIP-­ elicited PRL secretion in healthy individuals.167 There is no evidence that vasopressin stimulates PRL secretion. Kisspeptin stimulates prolactin secretion in animal paradigms, an effect that is mediated via the Kiss1 receptor.168 Further study is needed to examine whether kisspeptin stimulates prolactin secretion in humans. Prolactin-­releasing peptide (PrRP) is expressed in neurons in the paraventricular and supraoptic nucleus of the hypothalamus.169,170 In vitro, PrRP stimulates PRL secretion.170 However, its physiologic role with regard to the regulation of PRL secretion has not been elucidated.

PROLACTIN ACTIONS • P  rolactin exerts its actions by stimulating its receptor, a member of the cytokine receptor superfamily. • Prolactin has major physiologic effects on the breast and possibly other tissues. • Prolactin in excess causes hypogonadism, mediated primarily via hypothalamic actions but also exerts effects on the ovaries and testes.    In humans, PRL has a well-­ established physiologic role on the breast during pregnancy and the postpartum period, wherein it is essential for lactation to occur after delivery.171 In addition, PRL in excess suppresses the activity of the

CHAPTER 3  Prolactin in Human Reproduction

hypothalamic-­pituitary-­gonadal axis in the nonpregnant state.85 In animals, PRL has been implicated in many additional roles, including growth and development, effects on beta cell function, electrolyte transport, osmoregulation, and effects on the skin and cartilage, in addition to effects on behavior, immune function, and tumorigenesis.171,172 None of these additional actions have been adequately substantiated in humans. It is likely that the physiologic role of PRL is species-­specific.

Prolactin Receptor Prolactin exerts its actions by activating its cognate receptor, which is a member of class 1 (hematopoietic) cytokine receptor superfamily.173 In humans, the PRLR gene is located on chromosome 5 and contains 10 exons, which yield several diverse receptor isoforms as a result of alternative splicing of the primary transcript.174,175 These are broadly grouped into long, intermediate, and short isoforms, most of which possess the same extracellular and transmembrane domains but differ in the length of the intracellular domain.176 There is also a soluble isoform, which lacks both transmembrane and intracellular domains.173 The long prolactin receptor isoform is predominant in humans and is widely expressed in many cell types.173 The extracellular domain contains two disulfide bridges that are critical for PRL binding.177 The intracellular domain contains two conserved regions (Boxes 3.1 and 3.2), one of which (Box 3.1) is recognized by tyrosine kinases.178 The long isoform receptor is already dimerized before ligand binding occurs and can only initiate signaling upon PRL binding to the extracellular domain of the receptor homodimer.179 Signaling primarily involves activation of the Janus kinase 2 (JAK2)–signal transducer and activator of transcription 5 (STAT5) pathway.173 In addition, the mitogen-­activated protein kinase (MAPK) pathway is activated, and the phosphatidylinositol 3 kinase (PI3K) pathway is also recruited.171,180 Activation of these pathways results in altered transcription of target genes in the nucleus that mediates PRL actions.

Prolactin Effects on the Breast Prolactin has a critical role in breast development during pregnancy and milk synthesis postpartum as well as in pathologic states of prolactin excess.2,181 Of note, PRL stimulates alveolar tissue growth indirectly by stimulating, in synergy with progesterone, the synthesis of the receptor activator of NF-­ kappa B ligand (RANKL) and insulin-­ like growth factor-­ II (IGF-­II), both of which act in a paracrine manner to induce alveolar tissue growth.2,182,183 In addition to PRL, several other hormones, including placental lactogen, GH, insulin, cortisol, thyroxine, estrogen, and progesterone, have important roles in the development of breast tissue during pregnancy in preparation for lactation.184 Of note, high levels of estrogen prevent milk production during pregnancy.185 In contrast, some degree of estrogen priming of the breast is needed for lactation and this likely explains the infrequent occurrence of galactorrhea in postmenopausal women and men with hyperprolactinemia. After the placenta is delivered, the rapid decline in estrogen relieves its inhibitory effect on milk production.185 Postpartum, suppression of PRL secretion by bromocriptine leads to a rapid decline in milk production.186 Galactorrhea is defined as the secretion of milky fluid from one or both breasts in women who have either never been pregnant or have stopped nursing for over 12 months.187,188 Galactorrhea may rarely occur in men.188 Galactorrhea has been reported in 1%-­45% of women, depending on examination techniques and the patient population under study, and can be an important clue to the presence of PRL excess.187 Evaluation of PRL levels

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BOX 3.1  Causes of Hyperprolactinemia Physiologic Pregnancy Nursing Sleep Food consumption Coitus Stress Medications Antipsychotic agents (phenothiazines, haloperidol, risperidone, paliperidone, and others) Monoamine oxidase inhibitors Fluoxetine Opioids Cocaine Verapamil Alpha methyldopa Metoclopramide Domperidone Pituitary Prolactinomas Somatotropinomas and other cosecreting tumors Stalk effect in patients with nonfunctioning sellar and suprasellar masses Radiation therapy Stalk transection Systemic Primary hypothyroidism Renal failure Cirrhosis Adrenal insufficiency Polycystic ovary syndrome Spinal cord lesions Chest wall lesions Nipple stimulation Ectopic prolactin secretion Prolactin receptor gene mutations Idiopathic

and thyroid function is advisable in women with galactorrhea. However, only 28% of women with galactorrhea and regular menses have hyperprolactinemia.188 Indeed, women who have previously been pregnant can have persistent galactorrhea for several years in the absence of pituitary pathology.

Prolactin Effects on Gonadotropin Secretion In experimental animals, PRL appears to have an essential role in the regulation of the reproductive axis. Indeed, mice with targeted disruption of either the PRL gene or the PRL receptor gene have abnormal estrous cycles and are infertile.181,189 Whether physiologic PRL levels have an important role in regard to the regulation of the reproductive axis in humans is less clear. Healthy women who are administered bromocriptine to decrease PRL levels to ∼5 ng/mL have no change in gonadotropin pulsatility but exhibit higher estradiol levels in the late follicular phase and lower progesterone levels during the luteal phase of the menstrual cycle.190 Hyperprolactinemia leads to a marked decrease in gonadotropin secretion, including decreased LH pulse frequency and amplitude, as demonstrated by frequent serum sampling (Fig. 3.2).191,192 In menopausal women, hyperprolactinemia blunts gonadotropin levels, which are restored by bromocriptine therapy that lowers PRL levels.193

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Serum LH (mU/mL)

20

Before operation

10

0 0

Serum LH (mU/mL)

20

180

360

180

360

8 days after operation

10

0 0

Time (min) Fig. 3.2 Impaired luteinizing hormone (LH) pulsatile release was present in a hyperprolactinemic patient (top panel), which was restored after tumor resection (lower panel).  Secretory pulses are shown by arrows (Adapted from Stevenaert et al. Early normalization of luteinizing hormone pulsatility in women with microprolactinomas. J Clin Endocrinol Metab. 1986;62:1044–1047, with permission.)

Prolactin appears to exert direct effects on hypothalamic neurons to suppress GnRH secretion. In vitro, PRL leads to a decrease in GnRH secretion by hypothalamic neuronal cell lines.194 Hyperprolactinemic mice develop central hypogonadism, characterized by low gonadotropin and sex steroid levels.195 These animals have decreased kisspeptin 1 expression in the arcuate and anteroventral periventricular nucleus.195 These data are corroborated by studies in rats, which are also consistent with the hypothesis that PRL in excess blunts kisspeptin 1 expression in the hypothalamus.196 In hyperprolactinemic mice, administration of kisspeptin 1 restores the function of their reproductive axis, consistent with the hypothesis that kisspeptin 1-­expressing neurons in the hypothalamus have a major role in the pathogenesis of hyperprolactinemia-­induced hypogonadism.195 The role of kisspeptins as potential therapeutic agents in hyperprolactinemia-­ induced infertility requires study.197 In experimental animals, hyperprolactinemia blunts gonadotropin secretion after GnRH administration, suggesting that PRL may also directly influence gonadotroph function.198 However, the findings of similar studies in humans have been less consistent with observations made in animals.199 Hyperprolactinemic women also exhibit a lack of a positive effect of estradiol on gonadotropin secretion.200

Prolactin Effects on the Ovary In experimental animals, PRL has a role in maintaining the function of corpus luteum after ovulation.201 At physiologic levels, PRL stimulates the expression of 3 beta-­hydroxysteroid dehydrogenase (type 2) and has an important role in stimulating progesterone synthesis in human granulosa cells.202,203 However, studies in humans have not been conclusive with regard to the physi ologic role of PRL on ovarian function.

bromocriptine administration that suppresses PRL levels attenuates progesterone secretion and leads to shorter luteal phases in some, but not all, studies.204,205 At high levels, PRL inhibits aromatase expression in rat granulosa cells and decreases estradiol and progesterone synthesis in human ovaries.206–208 In aggregate, available data suggest that PRL excess suppresses the reproductive axis in women at several levels, leading to oligo-­amenorrhea. It should be noted that patients presenting with both amenorrhea and galactorrhea are likely to have hyperprolactinemia (75% of 471 patients in aggregated case series).188 Typically, these patients present with secondary, rather than primary, amenorrhea. Low libido and sexual dysfunction are also common in this group.209 On the other hand, young patients with hyperprolactinemia developing before the onset of puberty may present with primary amenorrhea.210 These younger patients appear to be less likely to have galactorrhea and may even present with pubertal delay as a result of estrogen deficiency, induced by PRL excess.210 Hyperprolactinemia appears to be common among women presenting with infertility, representing about one-­ third out of 367 women with infertility in aggregated case series.188 In a more recent study of women with infertility, the prevalence of hyperprolactinemia was 46%.211 Although most of these patients also have amenorrhea and galactorrhea, some patients manifest neither symptom but nevertheless have shorter luteal phases.212 Treatment of hyperprolactinemia in these patients increases progesterone synthesis during the luteal phase and improves fertility.212,213 Preovulatory, transient hyperprolactinemia in eumenorrheic women has also been associated with infertility.214 Hyperprolactinemia is variably present in women with polycystic ovary syndrome (PCOS) and it is presumed to be a consequence of sustained elevations in estradiol levels that stimulate PRL secretion, though this hypothesis is not universally accepted.215,216 In these patients, bromocriptine therapy that suppresses PRL levels may lead to a decrease in testosterone and LH levels and regular menstrual cycles.215,216

Prolactin Effects on the Testis Whether PRL at physiologic levels is relevant to the regulation of the reproductive axis in males has been a matter of debate. Male mice with targeted deletion of the PRL gene have normal testosterone levels despite decreased gonadotropin levels and are fully fertile.217 On the other hand, approximately 50% of male mice with targeted disruption of the PRL receptor gene have decreased fertility despite apparently normal spermatogenesis.181 In healthy men, bromocriptine administration for 8 weeks leads to a decrease in testosterone levels, both at baseline as well as after human chorionic gonadotropin (hCG) administration.218 In human sperm, PRL increases fructose utilization, glycolysis, and glucose oxidation.219,220 Available data suggest that PRL in excess suppresses the reproductive axis in men at several levels. In addition to decreased gonadotropin pulsatility, both baseline and hCG-­stimulated testosterone levels are often reduced in hyperprolactinemic men.221– 223 These patients frequently present with sexual dysfunction (low libido and erectile dysfunction) as a consequence of central hypogonadism.221–223 Up to 25% of men presenting with erectile dysfunction are hyperprolactinemic.224,225 It has been suggested that hyperprolactinemia per se may be associated with erectile dysfunction but the underlying mechanisms have not been elucidated.226 Gynecomastia and/or galactorrhea may develop in hyperprolactinemic men but are much less frequent than sexual dysfunction in this population.221–223 Infertility may also occur in hyperprolactinemic men.227 Up to 5% of men presenting with infertility have been reported to be hyperprolactinemic.227 Cabergoline therapy that suppresses PRL excess often leads to improvement

CHAPTER 3  Prolactin in Human Reproduction

in sexual function and recovery of the reproductive axis provided that pituitary gonadotrophs remain intact.228,229 Whether PRL normalization is essential for the recovery of erectile function is uncertain.221

Prolactin Effects on the Adrenal Cortex Hyperandrogenism of adrenal origin, based on the presence of elevated dehydroepiandrosterone sulfate (DHEA-­ S) levels, has been reported in some, but not all, studies of women with hyperprolactinemia.230,231 Elevation in testosterone levels may also occur in some patients as a consequence of conversion from androgenic precursors. Hirsutism may occur in women with elevated PRL levels but it is not clear if it correlates well with DHEA-­S levels.230,231 Bromocriptine therapy that suppresses PRL excess may lead to improvement in DHEA-­ S levels in women with hyperprolactinemia.212

Prolactin and the Skeleton Several lines of evidence suggest that abnormalities in PRL levels or action are often associated with abnormal bone metabolism. Mice with targeted deletion of the PRL receptor gene have low bone mineral density (BMD) and decreased bone formation.232 These animals also have low sex steroid levels, which may account for the low BMD rather than the absence of PRL action.232 In humans, PRL excess is associated with deleterious effects on bone. Women and men with hyperprolactinemia and hypogonadism have decreased BMD and increased risk of morphometric vertebral fractures.233–237 Dopamine agonist therapy that suppresses PRL excess leads to improvement in BMD.238 Of note, hyperprolactinemic women who have regular menstrual cycles have normal BMD.239 These data are most consistent with the hypothesis that PRL-­induced sex steroid deficiency, but not PRL excess per se, underlies the development of low BMD in hyperprolactinemic patients.239

PATHOLOGICAL STATES OF PROLACTIN DEFICIENCY AND EXCESS P •  rolactin deficiency generally signifies extensive pituitary failure. • Prolactin excess (hyperprolactinemia) can be caused by physiologic mechanisms, medications, pituitary and systemic etiologies, or can be idiopathic.

Prolactin Deficiency Prolactin deficiency may occur in patients with pituitary macroadenomas or other large sellar masses, wherein it is frequently associated with multiple additional pituitary hormone deficiencies.240 In these patients, the presence of PRL deficiency generally indicates advanced pituitary failure as a consequence of the destruction of pituitary tissue by tumoral hemorrhage or pituitary surgery.240 Prolactin deficiency may also occur in women with Sheehan’s syndrome (postpartum pituitary infarction in the setting of hemorrhagic shock as a consequence of obstetric complications).241 These patients classically present with failure of lactation and postpartum amenorrhea. Sheehan syndrome is currently uncommon in the US and other Western countries because of advances in obstetric care.241 Isolated, idiopathic PRL deficiency has been reported in 3 patients, who were unable to lactate, but suffered no other evident manifestations.242,243 In clinical studies, recombinant human PRL has been administered with success in some patients with PRL deficiency but is not currently FDA-­approved as replacement therapy.

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Hyperprolactinemia: Causes Hyperprolactinemia is relatively common in the general population (0.4%) and can be caused by diverse etiologies (Box 3.1).244–246

Physiological As already noted, PRL levels normally rise postprandially or during slow-­wave sleep. Acute stress (exercise, coitus, hypoglycemia, seizure, and others) may lead to a transient increase in PRL levels.101,103 Hyperprolactinemia is present during pregnancy and lactation, as already discussed, wherein it has a major physiologic role in breast development and milk production.53

Medications Older antipsychotic agents, including phenothiazines and butyrophenones, cause hyperprolactinemia by virtue of their dopamine antagonist properties.247 In these patients, hyperprolactinemia is generally mild to moderate (typically up to 100 ng/ mL) but can occasionally be more severe and may exceed 300 ng/ mL.248 Hyperprolactinemia resolves within 4 days after discontinuation of the offending agent.248 Some of the newer, “atypical” antipsychotic agents, including risperidone, paliperidone, and molindone, also cause substantial hyperprolactinemia, which may exceed 100 ng/mL.248 Other “atypical” antipsychotic agents, including olanzapine, quetiapine, aripiprazole, and ziprasidone, have a lesser effect on PRL levels as a consequence of weaker action on dopamine (D2) receptors and their partial dopamine agonist activity.248,249 As a corollary, the addition of low-­dose aripiprazole in patients with antipsychotic-­induced hyperprolactinemia reduces prolactin levels in patients taking risperidone, olanzapine, or other antipsychotic agents.250–252 Older, tricyclic antidepressants cause mild to moderate hyperprolactinemia in 25% of cases.248 Monoamine oxidase inhibitors have also been associated with mild hyperprolactinemia.253 Fluoxetine may raise PRL levels modestly, but these generally remain within the normal range.254 Other serotonin reuptake inhibitors, bupropion, nefazodone, trazodone, venlafaxine, and lithium do not cause hyperprolactinemia.248 Opioids and cocaine may also cause mild hyperprolactinemia, which may result in hypogonadism.248,255 Metoclopramide and domperidone inhibit dopamine receptors and commonly cause hyperprolactinemia and hypogonadism after chronic administration.248 There have been occasional case reports of hyperprolactinemia linked to the use of histamine (H2) receptor inhibitors, proton pump inhibitors, and protease inhibitors, but a causal relationship between these agents and PRL elevation has been disputed.248,256 Some older antihypertensive agents have been associated with hyperprolactinemia. Alpha methyldopa inhibits L-­aromatic amino acid decarboxylase, which converts L-­dopa to dopamine and acts as a false neurotransmitter interfering with dopamine secretion, thus causing hyperprolactinemia.248 Verapamil, but not other calcium channel antagonists (dihydropyridines and benzothiazepines), raises PRL levels by decreasing hypothalamic dopamine.257 Medications cannot be assumed to be the cause of hyperprolactinemia by default, although the timing between medication initiation and the onset of symptoms of hyperprolactinemia can be helpful. In addition, if serum PRL is known to be normal before the initiation of therapy with a medication associated with elevated PRL levels, then a cause-­and-­effect relationship between the medication in question and hyperprolactinemia can reasonably be inferred. However, a baseline PRL level is often not available in these patients. In that case, a different approach is required. If the potentially offending medication can be safely

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discontinued, then PRL levels should be checked 3 to 4 days later. If PRL levels normalize, then hyperprolactinemia can be attributed to the drug. If the medication cannot be safely discontinued, the patient should undergo pituitary imaging to detect a possible sellar mass (as detailed below).

Pituitary In addition to PRL-­secreting adenomas and PRL cosecreting tumors, which are discussed separately, virtually any mass lesion involving the pituitary and stalk may lead to hyperprolactinemia by interfering with dopamine outflow through the hypothalamic hypophyseal portal system into the anterior pituitary (“stalk effect”).245,258 Hyperprolactinemia is generally mild to moderate in patients with “stalk effect,” with PRL levels generally being below 150 ng/mL in most cases.245,258,259 Previous radiation therapy to the brain, delivered by linear accelerator, has been associated with mild hyperprolactinemia, presumably as a consequence of hypothalamic dysfunction.260 Similarly, proton beam radiation therapy to the base of the skull has often been associated with hyperprolactinemia.261

Systemic Primary hypothyroidism has been associated with mild hyperprolactinemia in up to a third of patients and resolves with levothyroxine replacement.262 It has been proposed that elevated TRH secretion and increased sensitivity of lactotroph cells to TRH underlie the increased PRL levels in these patients.262 In rare cases, primary adrenal insufficiency has been associated with mild hyperprolactinemia, which resolves after glucocorticoid replacement.263,264 Chronic kidney disease is associated with hyperprolactinemia, which becomes most common in patients with end-­stage renal disease requiring renal replacement therapies.265 In these patients, hyperprolactinemia appears to be a consequence of decreased PRL clearance and increased secretion and resolves after renal transplantation.265 Cirrhosis has been associated with hyperprolactinemia in 5% to 100% of patients in different series and has been postulated to occur as a consequence of decreased dopamine release in patients with hepatic encephalopathy.266 Nipple stimulation, chest wall lesions (burns, thoracic zoster, etc.) as well as spinal cord injuries have been associated with hyperprolactinemia, presumably as a consequence of neurogenic stimulation of PRL release.267 Very rarely, ectopic PRL secretion has been documented in isolated patients with uterine leiomyoma, uterine tumor resembling ovarian sex cord tumor, gonadoblastoma, pituitary tissue present within an ovarian teratoma, bronchogenic carcinoma, and renal cell carcinoma.268–272

Prolactin Receptor Gene Mutations In one report, three sisters were found to have hyperprolactinemia in the absence of a pituitary tumor or other demonstrable etiology.273 Two of these patients presented with oligomenorrhea and one had infertility. Of note, the propositus had given birth to four children but was treated with bromocriptine because of persistent galactorrhea postpartum.273 All three patients were found to have a heterozygous, germline mutation of the PRL receptor gene, resulting in a single amino acid substitution (His188Arg), which reduces signal transduction through the JAK-­STAT pathway.273 Another patient presented with absence of lactation after each of her two pregnancies and was found to have hyperprolactinemia in the absence of a sellar mass.274 Hyperprolactinemia was attributed to loss-­of-­function mutations of the PRL receptor gene. The patient was a compound heterozygote for two inactivating mutations of the PRL receptor gene and her unaffected parents were heterozygotes for these mutations.

Idiopathic In some patients with hyperprolactinemia, no evident etiology can be found. In such cases, hyperprolactinemia is termed idiopathic. The underlying mechanisms are likely heterogeneous and may reflect the presence of very small PRL-­secreting adenomas that are below the resolution of current pituitary imaging modalities in some cases, or underlying dysregulation of PRL secretion in other cases. On follow-­up, 23 out of 199 patients with idiopathic hyperprolactinemia developed pituitary microadenomas during a 2-­to 6-­year period.275,276 Of note, idiopathic hyperprolactinemia may resolve spontaneously in approximately one-­third of patients during follow-­up.277,278

Hyperprolactinemia: Diagnosis Patients with hyperprolactinemia require careful evaluation, including history and physical examination.85 Confirmation of PRL levels by repeat testing after overnight fasting is advisable, particularly in patients with mild hyperprolactinemia.85 Women of reproductive age should have a pregnancy test. Evaluation of kidney, liver, and thyroid function is also advisable. If no obvious cause is found to explain the presence of hyperprolactinemia, such as pregnancy or primary hypothyroidism, patients should undergo pituitary imaging by magnetic resonance imaging (MRI) using a pituitary protocol (or computed tomography [CT] if MRI is contraindicated, such as patients with some pacemakers).85,279 There is a good correlation between PRL levels and tumor size in patients with PRL-­secreting pituitary adenomas.85,245,280 As a corollary, patients with PRL-­ secreting macroadenomas generally have PRL levels over 250 ng/mL.85,245 In contrast, PRL levels tend to be lower (below 150 ng/mL) in patients with nonfunctioning sellar masses, causing “stalk effect.” As a word of caution, PRL levels should be rechecked in dilution in patients with very large sellar masses who have modest PRL elevations in order to detect a possible “hook effect” immunoassay artifact, which may occur in patients with exuberant PRL secretion (as already discussed).85,245 Patients with a microadenoma and modest PRL elevation may have either a functioning tumor (microprolactinoma) or a clinically nonfunctioning adenoma causing “stalk effect.”281 Regular pituitary imaging can be helpful in making this distinction, since a prolactinoma will generally shrink in size in response to dopamine agonist therapy, whereas a nonfunctioning adenoma will not get smaller, and may even enlarge, despite the institution of dopamine agonist therapy that normalizes PRL levels.

PROLACTIN-­SECRETING PITUITARY ADENOMAS (PROLACTINOMAS) P •  rolactinomas are common, benign pituitary tumors. • Clinical manifestations may include mass effect, hypopituitarism, and effects directly attributable to hyperprolactinemia. • Dopamine agonist therapy is generally the first-­line treatment for most patients who require therapy, with surgery and radiotherapy having important roles in selected cases.

Epidemiology and Natural History Prolactin-­secreting tumors represent the most common type of functioning pituitary adenoma.245 The incidence of these tumors was previously reported as being 10 cases per million per year and their prevalence is approaching 100 patients per million in the general population.282 However, more recent studies from several European countries, including Belgium, the United Kingdom, Finland, Iceland, and Switzerland, have suggested that

CHAPTER 3  Prolactin in Human Reproduction

the incidence and prevalence of these tumors are substantially higher (about five times) than previously reported.283–287 By definition, tumors below 10 mm in greatest diameter are termed microadenomas, whereas those that reach or exceed 10 mm are termed macroadenomas. This classification is clinically relevant, as larger tumors are more likely to cause mass effect or invade neighboring structures and have greater growth potential.245 Most prolactinomas are microadenomas. Of note, a larger proportion of macroadenomas occurs in men than women of reproductive age.245 However, macroadenomas have been reported to account for the large majority (94%) of PRL-­ secreting tumors that were diagnosed after menopause in a study of 17 women.288 A plethora of autopsy studies have identified the presence of clinically unsuspected pituitary adenomas in about 11% of subjects autopsied, with almost all (over 99%) of these tumors being microadenomas.289 These data are corroborated by the findings of imaging studies, which have identified the presence of incidental pituitary hypodensities consistent with microadenomas in 10% to 20% of study subjects.290,291 In autopsy studies, approximately 40% of adenomas examined by immunohistochemistry showed PRL immunostaining.292 Several studies have reported on the natural history of untreated microprolactinomas.275,293–295 Out of a total of 139 women followed for up to 8 years, tumor growth was detected in 9 patients (6.5%).275,293–295 These studies predated the use of MRI for pituitary imaging, but, nevertheless, do suggest that the growth potential of microprolactinomas is fairly low. Macroadenomas are proportionately more common in children or adolescents than in adults and are also more common in men than in women.245 Larger tumors tend to have a higher proliferation (Ki67) index.296 In some cases, tumor growth has been associated with estrogen administration.297 However, predictors and mechanisms of tumor growth remain poorly understood.296 It may be noted that most prolactinomas are benign, with frankly malignant tumors, which are, by definition, associated with intracranial or extracranial metastases, being exceedingly rare.245,298

Pathogenesis Prolactin-­secreting pituitary adenomas are monoclonal tumors.299 Factors involved in the pathogenesis of these neoplasms are unknown in most cases. Of note, mutations in the gene encoding the D2 dopamine receptor or the gene encoding the Gi2 alpha subunit, which couples the D2 receptor to adenyl cyclase, have been found in up to 15% of prolactinomas that are resistant to dopamine agonist therapy.300 Several germline, gain-­of-­function mutations in the gene encoding the PRL receptor, have been reported to be more frequent in patients with prolactinomas.301 Somatic mutations in several putative oncogenes or tumor suppressor genes, including ras, myc, Pit-­1, PROP-­1, c-­fos, MENIN, and others, have not been found in these tumors.299,302 Prolactin-­secreting pituitary adenomas develop in about 20% of patients with the multiple endocrine neoplasia 1 (MEN1) syndrome, caused by germline inactivating mutations of the MENIN gene, which encodes a tumor suppressor gene.303 Prolactinomas in these patients have been previously reported to behave more aggressively and be more resistant to medical therapy in comparison with sporadic tumors.304 On the other hand, more recent data in patients with the MEN1 syndrome suggest that prolactinomas generally respond well to medical therapy in this population.305 Parathyroid lesions are usually the presenting manifestation of MEN1 and are typically followed by the development of pituitary tumors.306 The coexistence of a prolactinoma (or another pituitary adenoma) and hypercalcemia secondary to primary hyperparathyroidism defines an index case for MEN1 and has been reported in approximately 14% of patients without known familial disease who were screened for hypercalcemia.

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one-­ third of these patients were also found to have gastrin-­ secreting enteropancreatic neuroendocrine tumors, which constitute another cardinal manifestation of MEN1.307 As a corollary, it is advisable to measure serum calcium levels in all patients with presumed prolactinomas. The syndrome of familial isolated pituitary adenoma (FIPA) has been the subject of substantial investigation in the recent past.308 In some of these patients, germline, inactivating mutations in the gene encoding the aryl hydrocarbon receptor-­interacting protein (AIP) have been identified and impart predisposition to pituitary tumor formation.309 This is transmitted as an autosomal dominant trait with incomplete penetrance and is associated with an increased risk of developing GH-­secreting tumors, prolactinomas, or cosecreting tumors.310–312 Pituitary adenomas present at a younger age in patients with AIP gene mutations and appear to be more aggressive and resistant to medical therapy.308 Approximately 10% of prolactinomas are resistant to dopamine agonist therapy.313 Decreased expression of D2 dopamine receptors, including the long dopamine receptor isoform, decreased D2 receptor density and dopamine binding sites on pituitary lactotrophs have been reported in these tumors and may underlie resistance to dopamine agonists.314,315 On the other hand, no association was observed between five D2 receptor gene polymorphisms and the response to cabergoline therapy in a study of 118 hyperprolactinemic patients.316 Decreased expression of transforming growth factor beta 1 (TGF beta1) and several components of its signaling cascade (Smad2 and Smad3) have been reported in tumors resistant to dopamine agonist therapy.317 Animal data suggest that TGF beta1 inhibits lactotroph proliferation and PRL secretion, raising the possibility that restoration of TGF beta1 activity might be efficacious as a therapy in patients with prolactinomas that are resistant to dopamine agonists.318

Pathology Prolactin-­ secreting pituitary adenomas are almost always benign.245 In rare cases, distant metastases within the cranium or in remote locations, such as liver, lungs, lymph nodes, or bone, may develop and are essential to define the presence of malignancy in these patients.298,299 Some prolactinomas may cosecrete other hormones, most often GH, but in some cases TSH, ACTH, or FSH.319–321 Acidophil stem cell adenomas often have oncocytic features and cosecrete PRL and GH.319,322 Symptoms attributable to hyperprolactinemia most often predominate over those related to GH excess in these patients.319,322

Clinical Manifestations In addition to symptoms and signs directly attributable to PRL excess, patients with PRL-­secreting macroadenomas may manifest symptoms related to mass effect or anterior hypopituitarism.245 Headache may occur as a result of pressure exerted on the dura and can be particularly severe and acute in the setting of pituitary apoplexy (characterized by sudden-­ onset neurologic and neuroendocrine impairments), generally occurring as a consequence of hemorrhage within an adenoma.323 Tumors extending superiorly into the suprasellar cistern may impinge upon the optic apparatus, causing a variety of visual field deficits.324 Bitemporal hemianopsia is typically noted in patients whose tumors compress the optic chiasm.324 Central scotomas may develop in patients with compression of one of the prechiasmatic optic nerves and homonymous hemianopsia may occur as a consequence of impingement onto one of the optic tracts.324 Rarely, large tumors may extend into the third ventricle, causing obstructive hydrocephalus.245,325 Ischemic stroke as a result of middle cerebral artery compression by a large tumor has been reported in a child with a large

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Tumors extending laterally often involve the cavernous sinus and may encase the intracavernous segment of the internal carotid artery but are very unlikely to cause arterial narrowing.327 Clinical manifestations attributable to compression of the cranial nerves (III, IV, V1, V2, or VI) coursing through the cavernous sinuses, namely ophthalmoplegia, facial pain, or numbness, are very uncommon in patients with pituitary adenomas, including prolactinomas.328 Nevertheless, such symptoms may occur in patients with pituitary apoplexy or occasionally in patients with large, aggressive adenomas.323 Rarely, large tumors may impinge upon the ipsilateral temporal lobe, causing complex partial seizures.325 Tumors extending inferiorly often erode into the sphenoid base and may involve the sphenoid sinus or even the posterior nasopharynx.329 These patients may rarely develop cerebrospinal fluid rhinorrhea in response to dopamine agonist therapy.330 In these cases, medical therapy causes dramatic tumor shrinkage, which unmasks gaps present in the bone and dura, leading to cerebrospinal fluid leakage.330 These patients typically require neurosurgical intervention in order to close these defects and mitigate the attendant risk of meningitis.330 Patients with macroadenomas may also develop anterior hypopituitarism and all require a thorough evaluation of anterior pituitary function.85,245 Of note, diabetes insipidus is extremely unlikely to develop in patients with pituitary adenomas in the absence of surgical intervention.331,332 As a corollary, patients who present with central diabetes insipidus and a sellar mass should be suspected of harboring a nonadenomatous lesion (such as a craniopharyngioma or metastasis or an infiltrative lesion) rather than a pituitary adenoma.331 Hyperprolactinemia is generally proportionate to tumor size in patients with prolactinomas.245 Women frequently present with galactorrhea, secondary amenorrhea, or infertility.333 In a total of 21 case series (1621 women), galactorrhea was present in 85% and oligomenorrhea in 93% of women.334 Men often present with low libido, erectile dysfunction, infertility, gynecomastia, or, uncommonly, galactorrhea.333 In a total of 16 case series (444 men), erectile dysfunction was present in 78% and galactorrhea was present in 11%.334 As already mentioned, men are more likely than women to harbor macroadenomas and are more prone to have invasive tumors.245,334–336 Whether this gender difference is only related to biologic factors, such as sex steroids, which may influence tumor behavior is debatable; delayed recognition of pituitary adenomas in men who may ignore their symptoms and attribute them to aging is possible in some cases.245,334 Children and adolescents may present with primary amenorrhea and delayed puberty.337,338 As already noted, younger patients are more likely to have macroadenomas, which are resistant to dopamine agonist therapy.339 Adolescents with hyperprolactinemia are at increased risk of developing low BMD, which is likely a consequence of hypogonadism occurring during a critical time of peak bone mass acquisition.340 An increase in all-­ cause mortality risk has been reported among patients with macroprolactinomas in comparison with the general population.341 There was no increase in mortality among patients with microprolactinomas in the same study. In addition, there was no association between the magnitude of hyperprolactinemia and mortality risk.341 It is conceivable that excess mortality can be accounted for by the presence of anterior hypopituitarism among some patients with macroprolactinomas. Further study is needed in order to elucidate this issue.

Management Overview The goals of therapy differ among patients with prolactino mas, depending on their symptoms, tumor size, presence of

hypogonadism, and patients’ interest in pregnancy.85,245 In patients with macroadenomas, prompt decrease in tumor size and long-­term tumor control, relief of mass effect, and restoration of pituitary function are all important goals of therapy. In addition, PRL normalization is desirable in all hyperprolactinemic patients who have hypogonadism or desire fertility in order to restore their gonadal function and optimize their fertility potential. Other possible indications for therapy include the presence of bothersome galactorrhea, gynecomastia, hirsutism, and acne, all of which may improve with PRL normalization.85,245 Medical therapy with a dopamine agonist represents an appropriate first-­line approach for most patients with prolactinomas who require treatment, including those with giant prolactinomas (≥4 cm in diameter).85,245,342 On the other hand, patients with microadenomas who have central hypogonadism, but are not seeking immediate fertility, can be treated with sex steroid hormone replacement therapy or an oral contraceptive instead of dopamine agonists, provided that their tumors remain stable during follow-­up. Studies in patients with idiopathic hyperprolactinemia or microprolactinomas have not shown a deleterious effect of oral contraceptives on PRL levels or tumor size.343,344 Asymptomatic patients with hyperprolactinemia and microadenomas who have normal gonadal function and are not seeking to improve fertility can be followed expectantly.85,245 In this group, it is advisable to monitor the patients’ PRL levels, periodically reassess their gonadal function and obtain follow-­up pituitary MRI examinations in order to detect possible tumor progression (reported to occur in about 7%–10% of patients over a 4–6 year interval, although not all changes were clinically significant) or the subsequent development of hypogonadism, either of which would require the institution of therapy.85,245 Although PRL levels and tumor size are generally well-­correlated in patients with prolactinomas, it should be noted that tumor growth has rarely been reported in the absence of a change in PRL levels.345,346 This observation suggests that periodic pituitary imaging is prudent in patients followed expectantly in order to detect this unusual possibility.

Medical Therapy Dopamine agonist therapy represents the cornerstone of management of most patients with prolactinomas who require therapy.245 Bromocriptine and cabergoline are both ergot alkaloid derivatives and are currently approved by the Food and Drug Administration (FDA) for use in patients with hyperprolactinemia and prolactinomas.245 Pergolide is no longer available in the US.245 Quinagolide, a nonergot compound, was never introduced in the US but is available for use in several other countries.334 Bromocriptine was the first drug in this class used to treat prolactinomas and hyperprolactinemia.347 Bromocriptine normalizes PRL levels in approximately 75% of patients and leads to the resumption of menses in the majority of treated women.347 In addition to inhibiting PRL secretion, bromocriptine inhibits DNA synthesis and cell proliferation, leading to a decrease in tumor size.348,349 Bromocriptine has been reported to induce autophagy and cell death in vitro.350 About 76% of patients treated with bromocriptine experience a decrease in tumor size over a treatment period of up to 10 years.334,347 An example of a tumor response to bromocriptine therapy is shown in Fig. 3.3. Tumor size reduction is variable in patients treated with bromocriptine. In approximately 40% of 112 patients reported in a total of 10 studies, there was a greater than 50% decrease in tumor size after the institution of bromocriptine therapy.334,347 Some tumors can completely disappear on follow-­up MRI examinations. In patients with macroadenomas impinging on the chiasm, an improvement in visual field deficits occurs in over 80% of treated patients and may be noticeable within 1 to 3 days Visible decreases

CHAPTER 3  Prolactin in Human Reproduction

Fig. 3.3 Sagittal (left, top, and bottom panels) and coronal (right, top, and bottom panels) magnetic resonance images of a patient with hyperprolactinemia and a pituitary macroadenoma, obtained before (top) and after (bottom) the institution of bromocriptine therapy.  There is substantial decrease in tumor size on medical therapy (From Molitch ME. Medical treatment of prolactinomas. Endocrinol Metab Clin North Am. 1999;28:143–169, with permission.)

in tumor size occur with a variable time course and can be observed within a period of several weeks to several months on therapy.351 Pituitary function may also improve in patients whose tumors shrink on bromocriptine therapy.352 The extent of tumor shrinkage and the magnitude of reduction in PRL levels are not well correlated in all patients.245,347 Some of the patients whose tumors do not respond adequately to bromocriptine therapy may be effectively treated with cabergoline (as will be detailed below).245,347,353 Bromocriptine therapy is usually initiated at a dose of 0.625 mg to 1.25 mg daily, taken in a single nightly dose to optimize tolerance, and is advanced every 3 to 7 days to a dose of 2.5 mg daily.245,347 Dose titration may occur in 4-­week intervals toward achieving normal PRL levels, provided that the medication remains well-­ tolerated. The medication is generally given in two or even three divided daily doses when the total daily dose exceeds 2.5 mg daily to improve effectiveness and tolerance; total daily doses exceeding 10 mg are rarely needed.245,347 Adverse effects associated with bromocriptine therapy most commonly include nausea and orthostatic dizziness.245,347 These can be minimized with gradual dose titration and intravaginal administration in some cases. In some patients, these adverse effects may improve over time. Less common adverse effects include headache, nasal congestion, constipation, digital vasospasm, vivid dreams, or nightmares.245,347 Pituitary tumor fibrosis may occur in some patients treated with bromocriptine and hinder subsequent attempts at transsphenoidal resection.354 Rarely, systemic fibrotic manifestations have been reported, including mediastinal and retroperitoneal fibrosis.347 The association between the use of some dopamine agonists and cardiac valvulopathy will be discussed below.355 Other adverse effects associated with dopamine agonist therapy include the development of psychosis or impulse control disorders, which resolve after medication discontinuation.356–360 Patients with a history of psychosis are, in general, not candidates for dopamine agonist therapy.

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can rarely occur even in patients without such a history. Medical therapy of prolactinomas in patients prescribed neuroleptics, which act as dopamine antagonists, should be carefully reviewed with the treating psychiatrist in order to avoid precipitating an increase in psychiatric symptoms. All patients receiving dopamine agonist therapy should be warned of the risk of psychiatric adverse effects, including impulse control disorders.361 Patients who develop impulse control disorders or psychosis should be withdrawn from dopamine agonists. Among patients whose tumors decrease in size in response to bromocriptine therapy, the medication dose can often be downtitrated, as the therapeutic benefits can generally be maintained on a lower dose.347 Eventually, some patients can be considered to be weaned off the medication after a treatment period of 2 to 3 years. Good candidates for dopamine agonist withdrawal are those who have maintained normoprolactinemia and have no visible tumor on follow-­up MRI examinations. Of note, recurrence of hyperprolactinemia may occur after bromocriptine withdrawal.347 About 20% to 50% of patients with microadenomas and 16% of patients with macroadenomas maintain normal PRL levels after medication withdrawal.362–364 As a corollary, patients who are withdrawn from therapy should be warned of the risk of recurrence and monitored with serial PRL levels and MRI examinations to detect possible recurrence, which should prompt the reinstitution of therapy. Cabergoline is another ergot alkaloid derivative with a very long half-­life after oral administration, which generally allows for infrequent dosing (once or twice a week).245 Cabergoline has a high affinity for D2 receptors in the pituitary and is very slowly eliminated as a consequence of its slow release from pituitary binding sites and extensive enterohepatic cycling.365 Cabergoline is more effective than bromocriptine in normalizing PRL levels and restoring gonadal function and is generally better tolerated.229,347,366 In one partially double-­blind randomized clinical trial of 459 women, most of whom had microprolactinomas, cabergoline therapy led to PRL normalization in 83% and restoration of regular menses in 72% of patients.366 In the same study, bromocriptine therapy led to PRL normalization in 59% and restoration of regular menses in 52% of patients.366 Similarly, cabergoline therapy leads to restoration of gonadal function in the large majority of hyperprolactinemic men.229 Cabergoline is also effective in reducing tumor size in most patients with prolactinomas.334,367–370 Patients who achieve more robust control of hyperprolactinemia on cabergoline therapy are likely to have a greater decrease in tumor size.371 In one study of hyperprolactinemic patients with macroadenomas who were treated with cabergoline therapy, 96% of 26 treatment-­ naive patients experienced a decrease in tumor diameter exceeding 50% over baseline.372 In the same study, 64% of 33 patients who were resistant to bromocriptine experienced a decrease in tumor diameter exceeding 50% over baseline after the institution of cabergoline therapy.372 Overall, available data indicate that many patients who do not respond to bromocriptine therapy can be treated effectively with cabergoline.245 Patients who have responded to cabergoline therapy only rarely experience tumor growth in the absence of recurrent, on-­therapy hyperprolactinemia (2 out of 115 patients in one study).373 Cabergoline therapy is usually initiated at a dose of 0.25 to 0.5 mg per week, taken at bedtime with a small snack, and is generally titrated every 4 to 6 weeks, aiming at restoring normoprolactinemia.245 More rapid dose titration can be appropriate in patients with large tumors causing mass effect. Most patients can be effectively treated with a total dose of 3.5 mg per week or less. About 10% of patients can be resistant to cabergoline (as well as bromocriptine), even in high doses, and may often require additional therapies, including pituitary surgery, radiation therapy, and/or temozolomide chemotherapy (as will subsequently be

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Several observational studies have reported a decrease in body mass index among hyperprolactinemic patients treated with cabergoline or bromocriptine.374 Uncontrolled, retrospectively collected data have suggested that cabergoline therapy in hyperprolactinemic patients may lead to a decrease in body weight and adiposity via a prolactin-­independent mechanism that involves an increase in resting energy expenditure.375 Randomized controlled trials are needed to establish the clinical significance of these observations. Cabergoline use has been associated with the same adverse effects noted among patients who are treated with bromocriptine, albeit less frequently.245 In a clinical trial of 459 hyperprolactinemic women, cabergoline therapy was discontinued in 3% of women because of adverse effects, whereas bromocriptine was discontinued in 12% of women because of adverse effects.366 Many patients who cannot tolerate bromocriptine because of nausea or dizziness can be treated safely with cabergoline, administered either orally or intravaginally.245,347 Indeed, published guidelines recommend using cabergoline over bromocriptine in most cases, based on greater efficacy and better tolerance of cabergoline therapy, except in the setting of preconception or pregnancy.85 Cardiac valvulopathy has been reported in some patients with Parkinson’s disease who were treated with very high doses of cabergoline (or pergolide, but not bromocriptine).376,377 In these patients, fibrotic lesions in the endocardium occur as a consequence of serotonin (5HT2B) receptor activation by cabergoline and can lead to clinically significant valvular regurgitation.355 A higher cumulative dose exposure is associated with a higher risk of valvulopathy.376,377 In contrast, patients treated with usual cabergoline doses (up to 2.0 mg/week) appear to be at low risk of valvulopathy.378–380 It appears sensible to inform patients as to available data and consider performing periodic echocardiographic monitoring on a case-­ by-­ case basis (particularly in patients who require larger than usual cabergoline doses).381 Among patients whose tumors decrease in size in response to cabergoline therapy, the medication dose can often be downtitrated, as the therapeutic benefits can generally be maintained on a lower dose.245,382 Cabergoline withdrawal can eventually be considered in patients who have responded well to medical therapy for at least 2 to 3 years, remain normoprolactinemic, and have no visible residual tumor on MRI.383–385 In a study of hyperprolactinemic patients who had achieved normoprolactinemia on cabergoline therapy, recurrence rates (5-­ year Kaplan-­ Meier estimates) for hyperprolactinemia after drug withdrawal were higher among patients with a visible residual tumor on MRI and were also higher among patients with macroadenomas than those with microadenomas.383,384 Among 105 patients with microadenomas, hyperprolactinemia recurred in 42% of patients with a visible residual tumor on MRI and 26% of patients without visible residual tumor at the time of medication withdrawal.383,384 Among 70 patients with macroadenomas, hyperprolactinemia recurred in 78% of patients with a visible residual tumor on MRI and 33% of patients without visible residual tumor.383,384 All patients who are eventually withdrawn from cabergoline therapy should be made aware of the risk of recurrence and be followed with serial PRL levels and pituitary MRI examinations.245 Pergolide is another dopamine agonist, which was previously used off-­label in the US to treat hyperprolactinemia.347 Pergolide appears to be comparable to bromocriptine with regard to efficacy and tolerability in patients with hyperprolactinemia.386 In high doses, pergolide was also used to treat Parkinson disease and was FDA-­approved for that indication. The medication was withdrawn from the US in 2007 because it was associated with cardiac valvulopathy when administered in high doses in patients with Parkinson disease.376,377

Quinagolide is a nonergot dopamine agonist that has been used to treat hyperprolactinemia in some countries.387,388 Approximately 50% of hyperprolactinemic patients who are resistant to bromocriptine can be treated effectively with quinagolide.387,388 In addition, some patients tolerate quinagolide better than bromocriptine.387,388 Quinagolide is not approved by the FDA and is not currently available in the US. First-­ generation somatostatin receptor ligands, including octreotide and lanreotide, are generally not effective in patients with prolactinomas. Pasireotide, a second-­generation somatostatin receptor ligand with expanded receptor isoform specificity, appears to be promising as a potential treatment for patients with resistant or aggressive prolactinomas expressing somatostatin receptors, based on case reports.389,390 More data are needed to confirm and extend these observations. Pasireotide is not approved by the FDA for use in patients with prolactinomas. Raloxifene, a selective estrogen receptor modulator, was reported to decrease serum prolactin levels (by ∼25%) in 10 out of 14 patients with prolactinomas, who had not shown an adequate response to dopamine agonist therapy.391 Larger studies are needed to establish the risks and benefits of raloxifene therapy in this population. Raloxifene is not FDA-­approved for use in patients with prolactinomas.

Surgery Pituitary surgery is typically performed transsphenoidally using an operating microscope or endoscope. Transsphenoidal pituitary surgery is generally considered second-­ line therapy in patients with prolactinomas.85,245,392 Patients who should be considered for surgery include those with macroadenomas that do not respond to dopamine agonist therapy, including patients with persistent chiasm compression or tumor growth during a trial of medical therapy.85,245,393 Some cystic prolactinomas are not responsive to medical therapy and should be considered for resection.394 Men are more likely than women to harbor tumors that are resistant to medical therapy.336 Patients with such tumors are also candidates for surgical intervention.85,245 In addition, patients presenting with pituitary apoplexy should be considered for surgery. Another indication for surgery is intolerance of dopamine agonist therapy.85,245,393 The rare development of cerebrospinal fluid leak, manifesting as rhinorrhea, or the occurrence of pituitary apoplexy on medical therapy (either of which may rarely occur in patients with prolactinomas being treated with dopamine agonists) are additional indications for surgical intervention.85,245 Patients with psychotic depression or schizophrenia are not good candidates for dopamine agonist therapy, as these medications can exacerbate psychosis.85,245 These patients should be considered for pituitary surgery, particularly if they have large tumors that are encroaching upon the optic apparatus. Among patients with microadenomas, individual preference and their desire to avoid medical therapy, if possible, is another possible indication for surgical intervention.85,245,395 However, patients with microprolactinomas opting for pituitary surgery should be referred only to expert pituitary neurosurgeons, who can achieve the best possible outcomes.359,396,397 Women who have macroadenomas with suprasellar extension and are seeking fertility should be considered for pituitary surgery in order to decrease their risk of tumor progression during subsequent pregnancy, particularly if a trial of dopamine agonist therapy fails to reduce tumor size.85,245 In addition, women with microadenomas who are seeking fertility and are resistant to the effects of dopamine agonist therapy are candidates for pituitary surgery.85,245 However, these patients may also be treated with ovulation induction in order to help them conceive. The likelihood of achieving complete resection depends on tumor size and location as well as the skill and experience of the

CHAPTER 3  Prolactin in Human Reproduction

neurosurgeon.398 In aggregated data from 50 case series, PRL normalization was reported in 75% out of 2137 patients with microadenomas and 34% of 2226 patients with macroadenomas.334 Recurrence of hyperprolactinemia was reported in 18% of patients with microadenomas and 23% of those with macroadenomas. Long-­term remission of hyperprolactinemia occurs in about 60% of patients with microadenomas but only approximately 25% of those with macroadenomas who underwent pituitary surgery.334 However, pituitary surgery is very effective in quickly relieving mass effect.399 Patients with visual deficits and large tumors that do not promptly respond to dopamine agonist therapy should be referred to an expert pituitary neurosurgeon. In these patients, tumor debulking and decompression of the optic apparatus generally lead to improved visual outcomes.245,399 Postoperatively, pituitary function improves in approximately one-­third of patients who had evidence of anterior hypopituitarism preoperatively.400 However, new deficits in pituitary function may also develop in up to 20% of patients after surgery.400 Permanent diabetes insipidus is uncommon when surgery is performed by an expert pituitary neurosurgeon.399 Other postoperative complications include epistaxis and cerebrospinal fluid rhinorrhea.399 Very uncommonly, tumor bed hemorrhage, meningitis, or stroke may occur. The perioperative mortality risk is about 0.2% to 0.5% and approaches zero in expert hands.399 Perioperative morbidity is also lower among patients operated on by more experienced neurosurgeons.398

Other Treatment Modalities (Radiation Therapy and Chemotherapy) In patients with prolactinomas, the role of radiation therapy (radiotherapy) is limited at present.85,245 Radiation therapy is very effective for tumor control and is advisable in patients with macroadenomas who require tumor control and cannot be adequately treated with dopamine agonist therapy and surgery.85,245 Of note, PRL normalization has been reported in only 35% of patients treated with a combination of surgery and radiation therapy and generally takes 5 to 15 years after radiotherapy to occur.334 Increasingly, radiation therapy is administered using stereotactic techniques that deliver either photons (Gamma knifeTM, CyberknifeTM) or protons selectively to the target.401 These newer modalities largely spare normal brain tissue from radiation exposure in comparison with older, conventional radiotherapy methods.401 Stereotactic radiation therapy can be administered in a single fraction (a technique known as “radiosurgery”) in appropriate candidates, who have smaller tumors that are distant from the optic apparatus.401 Based on historical comparisons, it has been suggested that radiosurgery is associated with a faster endocrine response than conventional radiation therapy.402,403 Anterior hypopituitarism is frequent after radiation therapy regardless of the modality used for administration.401–403 This complication occurs in about 40% of patients 5 years after radiation therapy and becomes even more prevalent long-­ term.401–403 As a corollary, all patients who receive radiation therapy to the sella require lifelong yearly reassessment of pituitary function and replacement of hormone deficiencies as needed. Other uncommon complications of radiation therapy include optic neuropathy or other cranial neuropathies.401 Infrequent complications in patients who received conventional radiation therapy include stroke, temporal lobe necrosis, or secondary tumor formation, which may occur several decades after radiation therapy.401 Whether stereotactic radiation therapy is associated with a lower risk of developing these very infrequent complications remains unknown. Temozolomide is an orally active alkylating agent, which has useful activity against many gliomas, including glioblastoma

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multiforme.404 Temozolomide exerts its effects by methylating DNA bases, resulting in DNA fragmentation and cell death.405–407 This drug has been helpful for tumor control in several patients with large, locally aggressive prolactinomas or the rare PRL-­ secreting carcinomas that continue to enlarge despite adequate dopamine agonist therapy, surgery, and radiation therapy.408,409 Up to 73% of patients with aggressive PRL-­ secreting adenomas appear to respond to temozolomide, using either tumor size or PRL levels as treatment endpoints.405–407 Expression of the DNA repair enzyme O6-­methylguanine-­DNA methyl-­transferase (MGMT) in the tumor is inversely correlated with response rates to temozolomide in several, but not all, studies.405–407,410 Lack of treatment response after three cycles of temozolomide therapy predicts resistance to this agent in patients with aggressive pituitary adenomas (including prolactinomas).407 Escape from the salutary effects of temozolomide on tumor size may occur. Temozolomide is not approved by the FDA for use in patients with prolactinomas. Everolimus is an inhibitor of the mammalian target of rapamycin (mTOR), which might be of therapeutic benefit in patients with prolactinomas that are resistant to dopamine agonists, based on a case report.411 More data are needed to establish the risks and benefits of everolimus therapy in patients with prolactinomas. Everolimus is not approved by the FDA for use in this population. Lapatinib, an EGF receptor/ErbB-­2 tyrosine kinase inhibitor, was reported to decrease PRL levels and tumor size in two patients with prolactinomas that were resistant to dopamine agonist therapy.412 In a phase IIa study of four patients, lapatinib therapy led to tumor stability in three patients with aggressive prolactinomas treated for 6 months.413 However, one patient with a prolactin-­secreting carcinoma showed disease progression after 3 months in this study.413 More studies are needed to characterize the effects of lapatinib in patients with aggressive prolactinomas. Lapatinib is not FDA-­approved for use in patients with prolactinomas.

PREGNANCY AND PROLACTINOMAS • P  rolactin normalization improves fertility in hyperprolactinemic patients. • In contrast to microprolactinomas, macroprolactinomas have a substantial growth potential during pregnancy. • Use of bromocriptine in the preconception period appears to be safe for the fetus and the pregnancy. • Cabergoline also appears to be safe in the preconception setting but pertinent data are more limited.

Prolactin-­Secreting Pituitary Adenomas During Preconception and Pregnancy If left untreated, hyperprolactinemia often decreases fertility potential (as already reviewed).188,212,245 In these women, PRL normalization generally improves their likelihood of conceiving.188,212,245 As a corollary, hyperprolactinemia is an indication for dopamine agonist therapy in all women presenting with subfertility.414 In these patients who are treated in the preconception setting, bromocriptine is considered the drug of choice (as will subsequently be detailed).245 Some prolactinomas may enlarge during gestation as a consequence of tumor exposure to the estrogenic milieu of pregnancy or withdrawal of previous dopamine agonist therapy (Fig. 3.4).415 As already discussed, normal lactotroph hyperplasia is physiologic during pregnancy and regresses within several months postpartum.416 In women with prolactinomas, the risk of tumor growth during pregnancy is higher among those with macroadenomas than women with microadenomas (Table 3.1).415 In aggregate, the risk of clinically significant tumor growth during

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Nursing has not been associated with an increase in size of prolactinomas or a higher risk of persistent hyperprolactinemia after lactation has ended.420–422 In the absence of a preexisting mass effect exerted by a growing adenoma, women should be encouraged to nurse. Dopamine agonist therapy will suppress milk output and the drug is present in breast milk if lactation is not completed suppressed. As a corollary, dopamine agonists should be avoided in this setting in the absence of compelling indications, including mass effect as a consequence of a growing tumor.

Effects of Dopamine Agonists During Preconception and Pregnancy

Fig. 3.4 Coronal (left, top, and bottom panels) and sagittal (right, top, and bottom panels) magnetic resonance images of a patient with hyperprolactinemia and a pituitary adenoma, obtained before pregnancy (top) and during the third trimester of pregnancy (bottom).  There was evident tumor growth during pregnancy in this patient, who reported persistent headache (From Molitch ME. Medical treatment of prolactinomas. Endocrinol Metab Clin North Am. 1999;28:143–169, with permission.)

TABLE 3.1  Outcomes of Women With Prolactinomas During Pregnancy, Stratified by Tumor Size at Baseline and Pre-­Pregnancy Management Tumor Size Before Pregnancy

Symptomatic Pre-­Pregnancy Tumor Growth Number of Surgery and/or During Women Radiation Therapy Gestation

Microadenoma 800 Macroadenoma 288 Macroadenoma 148

No No Yes

20 (2.5%) 52 (18.0%) 7 (4.7%)

(Data extracted from: Molitch ME. Prolactinoma in pregnancy. Best Prac Res Clin Endocrinol Metab. 2011;25:885–896; Rastogi A, Bhadada SK, Bhansali A. Pregnancy and tumor outcomes in infertile women with macroprolactinoma on cabergoline therapy. Gynecol Endocrinol. 2017;33:270–273; Karaca Z, Yarman S, Ozbas I, et al. How does pregnancy affect the patients with pituitary adenomas: a study on 113 pregnancies from Turkey. J Endocrinol Invest. 2018;41:129–141.)

pregnancy, causing mass effect, was 2.5% (20 out of 800 women) in patients with microprolactinomas.415,417–419 In several of these patients, bromocriptine therapy during pregnancy resulted in prompt symptomatic relief.415 In contrast, the risk of clinically significant tumor growth during pregnancy was 18.0% (52 out of 288 women) in patients with macroprolactinomas who did not undergo pituitary surgery or radiation therapy before conception.415,417–419 However, the risk of tumor growth during gestation was 4.7% (7 women) among 148 patients with macroadenomas who underwent surgery or radiation therapy before conceiving.415,417–419 As a corollary, medical therapy with bromocriptine is advisable before conception in order to reduce the size of macroadenomas before pregnancy.245,415 If an adequate tumor response cannot be achieved through medical therapy alone, then pituitary surgery should be considered in order to debulk the tumor and decrease the risk of adenoma growth dur ing the subsequent pregnancy.

No clinical trials of dopamine agonist therapy have been conducted during preconception or pregnancy. However, safety data have been published on women who conceived while taking dopamine agonist therapy.245,415 There are more limited data on women who were treated with a dopamine agonist throughout pregnancy. Bromocriptine use during preconception and very early pregnancy was not associated with an increase in congenital malformations, ectopic pregnancies, spontaneous abortions, or preterm deliveries, based on data from over 6000 pregnancies (Table 3.2).415,423,424 Children whose mothers received bromocriptine in the preconception setting have not been found to be at increased risk of developmental abnormalities during childhood.425 Data in about 100 women who were treated with bromocriptine for a longer time during pregnancy also suggest that bromocriptine use appears safe during gestation.415,426 However, it should be recognized that safety data in women treated with bromocriptine throughout pregnancy remain limited and therefore any conclusions regarding medication safety in this setting have to be made with caution. Cabergoline use during preconception and very early pregnancy was not associated with harm to the fetus or the pregnancy, based on data from about 1000 pregnancies.415,417–419 Children whose mothers were treated with cabergoline during preconception were not found to be at increased risk of developmental abnormalities.427–429 However, the safety database in patients treated with cabergoline in the preconception setting is smaller, comprising about 1000 patients in total.415,417,419,430–432 As a corollary, bromocriptine is the dopamine agonist of choice in hyperprolactinemic women during preconception, but cabergoline can be used if bromocriptine is not effective or tolerated. In contrast to previously reported findings, a retrospective study of 57,408 women from the EFEMERIS database, 183 of whom were treated with bromocriptine or cabergoline at some point during pregnancy, reported an increased risk of miscarriage and prematurity in association with dopamine agonist therapy but no significant difference in congenital malformations or infants’ psychomotor development.433 The significance of these observations remains incompletely understood. Safety data on pergolide and quinagolide have suggested the possibility of an increased risk of fetal harm in association with the use of these medications in the preconception period.415,434,435 Based on available data, it would be prudent to avoid these two medications in women wishing to conceive.

Management of Patients With Prolactinomas During Preconception and Pregnancy Based on our understanding of the pathophysiologic role of PRL excess, it would be advisable to treat all hyperprolactinemic women planning to conceive, regardless of whether they have regular menses or not.245,415 Bromocriptine is the drug of choice in this setting in light of its larger safety database in comparison with that of cabergoline. However, cabergoline also

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TABLE 3.2  Safety Data on Pregnancy Outcomes After Exposure to Bromocriptine or Cabergoline During the Preconception Period. After Bromocriptine Therapy N (%) After Cabergoline Therapy N (%) Number of pregnancies Miscarriages (spontaneous abortions) Ectopic pregnancies Terminations Hydatidiform moles Number of deliveries of known duration Deliveries at term (≥37 weeks) Preterm deliveries (COX-­1 mice (COX-­2 knocked in under the control of the COX-­1 promoter) can be corrected by COX-­2 exchange in COX-­2>COX-­1 mice, with normal term pregnancy, gestation length and litter size. In contrast, loss of native COX-­ 2 in COX-­ 1>COX-­ 2 mice results in severely impaired reproductive function.306 COX-­ 2, but not COX-­ 1, is induced during inflammation-­ mediated preterm labor elicited by lipopolysaccharide (LPS) administration.307 In a murine model, the COX-­2 selective inhibitor SC-­236 was more effective than the COX-­1 selective inhibitor SC-­560 in stopping LPS prompted preterm labor and increasing uterine PG synthesis.307 Furthermore, COX-­1 deficient mice, which show a delay in the onset of term labor, exhibit no delay in the onset of preterm labor after LPS treatment.307 Although it is likely that COX-­2-­derived PGF2α acts as a luteolysin in the mouse model of LPS-­induced preterm labor, it is not known whether the uterotonic effect of PGF2α and other COX-­2-­derived PGs may also play a role in this setting. The latter possibility may be particularly relevant to human preterm labor because indomethacin, an inhibitor of both COX-­1 and 2, has been used successfully in treating human preterm

CHAPTER 4  Steroid Hormones and Other Lipid Molecules Involved in Human Reproduction

labor when given systemically and when delivered locally through the vaginal route.308,309 The clinical utility of indomethacin as a tocolytic agent, however, is tempered by concerns over fetal and neonatal complications, such as the constriction of the DA.308,309 The adverse effects associated with indomethacin use in preterm labor have been mainly linked to inhibition of COX-­2 based on mouse studies using selective COX-­1 and COX-­2 inhibitors.310,311 COX-­1 selective inhibitors, should they be developed clinically, would be expected to produce fewer adverse effects than traditional NSAIDs when used as tocolytic agents.309-­311 Studies in animals and pregnant women comparing indomethacin and coxibs produced mixed results when fetal ductus blood flow was used as a measure of potential adverse effects.309

Eicosanoids and Ductus Arteriosus Remodeling As mentioned earlier in the chapter, one of the complications of using NSAIDs such as indomethacin to treat preterm labor is the induction of premature closure of the fetal DA.250,312 The DA is a large fetal vessel that shunts deoxygenated blood away from the pulmonary circulation to the descending aorta and to the umbilicoplacental circulation, where oxygenation occurs. In neonates, rapid remodeling of the DA leads to its closure after adaptation of spontaneous breathing in newborn infants. Although patency of the DA in utero is essential for proper fetal growth and development, failure of the DA to close after birth, called persistent patent DA, compromises postnatal health by causing circulatory complications such as pulmonary hypertension and congestive heart failure.312 Prostaglandins are intimately involved in DA function and its perinatal remodeling, and details of their mechanism of action are emerging. The finding that indomethacin administration induces premature DA closure in fetuses.250,312 suggested that fetal PGs are essential for maintaining DA patency. However, mice lacking both COX-­1 and COX-­2 are unable to make any PGs and die postnatally with patent DA,313,314 indicating that fetal-­derived PGs are not necessary for maintaining DA patency in utero. Such PGs do, however, play an indispensable role in DA remodeling after birth. How can this phenotype be reconciled with the results of pharmacologic inhibition studies? Researchers have proposed that PGE2 in the fetal circulation (supplied in part by the placenta) maintains dilation of the DA in utero and that COX-­2 in the DA produces constrictor PGs that are important for DA contraction after birth.314 Thus, indomethacin-­induced premature DA closure in fetuses may reflect inhibition of dilatory PGE2 synthesis in the placenta without sufficient inhibition of ductal COX-­2 to attenuate DA contraction. Although COX-­1 deficiency alone does not affect perinatal DA remodeling, it was found to exacerbate the phenotype of patent DA in the background of COX-­2 deficiency.314 In other studies, mice deficient in the smooth muscle relaxant receptor EP4315,316 or the PGE2-­metabolizing enzyme 15-­OH-­PGDH317 also fail to survive postnatally due to patent DA. EP4 expression in DA and 15-­OH-­PGDH expression in the fetal lung increase dramatically just before birth, supporting the conclusion that these proteins play an important role in perinatal DA remodeling. The phenotypes of EP4 deficient and 15-­OH-­PGDH deficient mice, together with the results of pharmacologic studies, support alternative models for the DA remodeling process.316-­323 Signaling through EP4 plays two essential roles in DA patency and remodeling, namely vascular dilation and intimal cushion formation (ICF). The vascular smooth muscle-­relaxing ability of PGE2 acting via EP4 maintains patency during fetal life. When PGE2 levels plummet at birth due to rapid and efficient metabolism by late gestation induced 15-OH-PGDH activity, the vaso dilatory role is curtailed, and functional DA closure is triggered

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rapidly by an increase in oxygen tension. ICF occludes the vascular lumen and results in permanent closure after birth. ICF within the DA is a result of an increase in vascular smooth muscle cell migration and proliferation and the production of hyaluronic acid (HA) under the endothelial layer, along with decreased elastin fiber assembly. PGE2 via EP4 signaling during late gestation controls a gene, HAS2, which regulates HA synthesis and ICF.323

Eicosanoids and Preeclampsia The possible involvement of PGs in preeclampsia and the potential therapeutic efficacy of low-­dose aspirin prophylactic treatment has received considerable attention. Preeclampsia occurs in 10% of pregnancies and is recognized as a prevalent source of risk to both mother and fetus. Although the exact cause of the disease is unknown, dysregulated production of PGI2 and TxA2 has been postulated as one of many potential etiologic factors.324 Decreased urinary PGI2 metabolites precede the development of preeclampsia,325 and increased TxA2 metabolite excretion occurs in patients with severe preeclampsia.327-­328 Such changes may predispose to vasoconstriction of small arteries, activation of platelets, and uteroplacental insufficiency—clinical outcomes that are associated with PIH and preeclampsia. In a murine model of enhanced TXA2 activity, transgenic overexpression of TP in the vasculature results in intrauterine growth retardation that is rescued by timed suppression of TXA2 synthesis with indomethacin.329 Many clinical trials have been conducted to evaluate the use of low-­ dose aspirin in the prevention of preeclampsia. Some randomized studies indicated a beneficial effect for women at increased risk, other trials did not show a positive effect of low-­ dose aspirin use on reducing the incidence of preeclampsia or on improving perinatal outcomes in pregnant women at high risk for preeclampsia.330-­333 Based on a recent evaluation of clinical trials, the US Preventive Task Force recommended daily low-­dose aspirin for women at high risk for preeclampsia.334,335

Eicosanoids in Male Reproduction Although PGs are critically involved in multiple steps of female reproduction, as previously discussed, their physiologic role in male reproduction is not well understood—though it appears to be less remarkable, based on gene knockout studies. No discernible male reproductive phenotype has been noted in any of the COX-­deficient or prostanoid receptor-­deficient mice.210,211 On the other hand, a number of studies have described androgen-­dependent regulation and distinctive tissue distribution patterns of COX-­1 and COX-­2 enzymes, as well as PG-­ synthesizing enzymes, in the male reproductive tracts of rodents and humans.336-­339 Specific functions have also been documented for COX-­2-­derived PGD2 and PGE2 in mediating cytokine production in Leydig cells and in regulating apoptosis in the rat epididymis.340,341 Although they are not definitive, such studies suggest that PGs may be synthesized and functional in a regulated manner in the male reproductive organs, even though their physiologic roles in these sites are dispensable. In the future, a better understanding of the eicosanoid network in the male reproductive system may lead to new PG-­based therapies or provide mechanistic insight into existing therapies. For example, a highly specific expression of COX-­2 has been found in the distal end of the rat vas deferens.336 Because the distal vas comprises an extensive submucosal venous plexus connected to the penile corpora cavernosa, PGs from the vas may play a role in erection.336 In this context, it is notable that intracavernosal injection of PGE1 has been used clinically as an effective therapy for erectile dysfunction in men.342,343 While PGs were originally isolated in large amounts from male seminal vesicles, their role at this site has remained enigmatic. The recent finding that ovarian

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PART I  The Fundamentals of Reproduction

PGs may act as sperm guidance factors could indicate a synergistic role with male accessory gland-­derived PGs.344 Furthermore, PGE2 may protect sperm from phagocytosis by neutrophils and inhibit pro-­inflammatory cytokine formation, enhancing sperm survival and improving the motility and binding capacity of sperm in the oviduct, which could increase the fertilization rate.345

OTHER LIPID MEDIATORS: LYSOPHOSPHATIDIC ACID AND SPHINGOSINE-­1-­PHOSPHATE Besides eicosanoids, other lipid mediators have been implicated in reproductive function, such as lysophosphatidic acid (LPA) and sphingosine-­1-­phosphate (S1P).346,347 LPA is a lipid-­signaling molecule and an intermediate in the de novo biosynthetic pathway of phospholipids consisting of a glycerol backbone, a phosphate head group, and a long-­chain fatty acid (usually oleic acid or palmitic acid), most commonly in acyl linkage (see Fig. 4.16).348 LPA generation is complex, proceeding by at least two pathways: conversion from lysophospholipids or via phosphatidic acid. Several phospholipase activities are necessary, including phospholipase A1 (PLA1)/PLA2 plus lysophospholipase D (lysoPLD) and phospholipase D(PLD). Additional extracellular phospholipases, such as secretory PLA2 (sPLA2-­IIA), membrane-­associated PA-­selective PLA1 (mPA-­ PLA1), and lecithin–cholesterol acyltransferase (LCAT), can also be involved.349 LPA signaling is mediated primarily by six members of the GPCR family currently referred to as LPA1, LPA2, LPA3, LPA4, LPA5, and LPA6.350 The lipid mediator S1P is a bioactive sphingolipid derived from the abundant phospholipid sphingomyelin. Sphingomyelinases generate ceramide, which is cleaved by ceramidases to sphingosine, followed by phosphorylation by sphingosine kinases to S1P (Fig. 4.22).351 S1P signals extracellularly via transmembrane receptors but also has intracellular targets.352,353 There are five S1P receptors, also members of the GPCR family, referred to as S1P1, S1P2, S1P3, S1P4, and S1P5.354 S1P stimulation of these

receptors elicits a battery of downstream effects, including inhibition of cAMP, and activation of mitogen-­ activated protein kinases, phospholipase C, and PI3 kinase to evoke a broad range of cellular activities.354

LPA and S1P in Reproductive Function LPA and S1P are two lysophospholipids that play important roles in reproduction acting via their respective GPCRs.355 Most of the LPA receptor genes have been disrupted in mice, with some affecting reproductive function. LPA3 (also known as Edg7) exhibits a female reproductive phenotype.356,357 LPA3 mRNA has been detected in the oviduct, placenta, and uterus, but not in ovary and oocytes, with expression highest early in pregnancy, at approximately embryonic day 3.5 in mice. LPA3 is regulated positively by progesterone and negatively by estrogen.356,357 LPA3-­ deficient females produce small litters and show a prolongation of pregnancy by approximately 1.5 days (normal gestation, 19.5 days).356 The mice show no obvious defects in ovulation, ovum transport, or blastocyst development. However, the defects are related to delayed implantation and altered positioning or crowding of embryos, which leads to delayed embryonic development and death, accounting for the reduced litter size. The observed phenotypes are the result of maternal LPA3 signaling and are not due to embryo LPA3 signaling.356 Strikingly, the phenotypes are similar to those seen with cPLA2 female knockout mice and rodents treated with indomethacin.358-­360 LPA3-­deficient female uteri had markedly reduced COX-­2 expression and PGE2/PGI2 levels at E3.5, thus linking LPA signaling to PG biosynthesis and fertility control.356 Overall, this particular study raises speculation that therapeutic manipulation of LPA3 signaling could influence the low implantation rate, which is a major drawback during infertility treatments using assisted reproductive technologies. LPA1, LPA2, and LPA3 are differentially expressed in testes with the latter two in the basal regions of seminiferous tubules, primarily in immature germ cells (spermatogonia and spermatocytes) and LPA1 showing stage-­specific expression in germ OH O O C

Sphingomyelin

NH2+

P

CH3 O

O−

CH2

CH2

N+

CH3

CH3

O Sphingomyelinase OH OH C

Ceramide

NH2+

O Ceramidase OH OH

Sphingosine

NH2 Sphingosine kinase + ATP OH

O O

Sphingosine 1-phosphate

NH2

P

OH

O−

Fig. 4.22 Structures and biosynthesis of a representative sphingosine-­1-­phosphate (S1P) lipid mediator from sphingomyelin.  Sphingomyelinase removes the phosphorylcholine head group to yield ceramide. Ceramidase cleaves the amide bond removing one aliphatic chain to generate sphingosine. In the presence of adenosine triphosphate

CHAPTER 4  Steroid Hormones and Other Lipid Molecules Involved in Human Reproduction

cells.361 Triple knockout mice exhibit male reproductive defects including alterations in mating behavior and heterogeneic spermatogenic disruption resulting in sterility.361 Many questions remain to be answered on the mechanisms for these effects. Of the “newer” LPA receptors LPA4, LPA5, and LPA6, there is no evidence for the involvement of these signaling pathways in reproductive function. LPA4 has been detected in ovaries, uterus, and placenta, but only one of three research groups reported a reduced litter size in mice deficient for LPA4, and this was apparently attributable to other nonreproductive mechanisms.362 Clues to the roles of S1P in reproductive function have come from various sources. Based on expression studies of the five receptor subtypes that bind S1P, only three (S1P1, S1P2, and S1P3) show widespread distribution in mice, with expression in gonadal tissues and in the uterus during decidualization.354,363 S1P1 and S1P2 colocalize with COX-­2 at the maternal/fetal interface throughout pregnancy, suggesting a link between sphingolipid and PG signaling and indicating that S1P coordinates uterine mesometrial angiogenesis during implantation.363 Because S1P1-­deficient mice die in mid-­embryogenesis as a result of complications of vasculogenesis, it has not been possible to decipher the role of this particular signaling pathway in reproductive function.354 Both S1P2-­and S1P3-­null mice show no obvious phenotypes with the exception of slightly smaller litter sizes.350,354 However, when both receptors are knocked out, there is infertility.350 The specific roles of each S1P receptor subtype in female reproductive function will require more study. The role of S1P in the preservation of female fertility is being recognized.364 Programmed cell death (apoptosis) is an established paradigm in the mammalian female germline. Ceramide generated by membrane cleavage of sphingomyelin by sphingomyelinase or via de novo biosynthesis by ceramide synthase is translocated from cumulus cells to adjacent oocytes to induce germ cell apoptosis.365 This is prevented by S1P, a ceramide metabolite within the same pathway, or by acid sphingomyelinase (ASM) deficiency. The therapeutic management of infertility by S1P in premature menopause and in female patients with cancer appears promising but awaits further study.364

109

Sphingolipids also seem to play a role in male germ cell apoptosis.366-­367 Ceramide induces an early apoptotic pathway event in male germ cells that is partially suppressed by S1P. However, although the maintenance of normal sphingomyelin levels in testes and normal sperm motility is dependent on ASM, testicular ceramide production and the ability of germ cells to undergo apoptosis do not require ASM. TOP REFERENCES

Auchus ML, Auchus RJ. Human steroid biosynthesis for the oncologist. J Investig Med. 2012;60(2):495–503. Baird DT, Horton R, Longcope C, et al. Steroid dynamics under steady-­ state conditions. Recent Prog Horm Res. 1969;25:611–664. Funk CD. Prostaglandins and leukotrienes: advances in eicosanoid biology. Science. 2001;294:1871–1875. Guo L, Ou X, Li H, et al. Roles of sphingosine-­1-­phosphate in reproduction. Reprod Sci. 2014;21(5):550–554. Hammond GL. Diverse roles for sex hormone-­binding globulin in reproduction. Biol Reprod. 2011;85(3):431–441. Henry HL. Regulation of vitamin D metabolism. Best Pract Res Clin Endocrinol Metab. 2011;25(4):531–541. King SR, Lavoie HA. Gonadal transactivation of STARD1, CYP11A1 and HSD3B. Front Biosci. 2012;1(17):824–846. Miller WL. Disorders in the initial steps of steroid hormone synthesis. J Steroid Biochem Mol Biol. 2017;165:18–37. Miller WL, Auchus RJ. The molecular biology, biochemistry, and physiology of human steroidogenesis and its disorders. Endocr Rev. 2011;32(1):81–151. Penning TM. Human hydroxysteroid dehydrogenases and pre-­receptor regulation: insights into inhibitor design and evaluation. J Steroid Biochem Mol Biol. 2011;125(1-­2):46–56. Sugimoto Y, Inazumi T, Tsuchiya S. Roles of prostaglandin receptors in female reproduction. J Biochem. 2015;157(2):73–80. Turcu AF, Rege J, Auchus RJ, Rainey WE. 11-Oxygenated andro­gens in health and disease. Nat Rev Endocrinol. 2020;16(5):284–296. Ye X. Lysophospholipid signaling in the function and pathology of the reproductive system. Hum Reprod Update. 2008;14(5):519–536. Ye J, DeBose-­Boyd RA. Regulation of cholesterol and fatty acid synthesis. Cold Spring Harb Perspect Biol. 2011;3(7):a004754.

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REFERENCES

1. Umetani M, Shaul PW. 27-­Hydroxycholesterol: the first identified endogenous SERM. Trends Endocrinol Metab. 2011;22(4):130–135. 2. Nixon M, Upreti R, Andrew R. 5α-­Reduced glucocorticoids: a story of natural selection. J Endocrinol. 2012;212(2):111–127. 3. Committee on the clinical utility of treating patients with compounded bioidentical hormone replacement therapy, board on health Sciences Policy, health and medicine Division, National Academies of Sciences, engineering, and medicine. In: Mattison DR, Parker RM, Jackson LM, eds. The Clinical Utility of Compounded Bioidentical Hormone Therapy: A Review of Safety, Effectiveness, and Use. National Academies Press; 2020. 4. Hu J, Zhang Z, Shen WJ, et al. Cellular cholesterol delivery, intracellular processing and utilization for biosynthesis of steroid hormones. Nutr Metab (Lond). 2010;1(7):47. 5. Rainey WE. Adrenal zonation: clues from 11β-­hydroxylase and aldosterone synthase. Mol Cell Endocrinol. 1999;151(1–2):151–160. 6. Miller WL. Disorders in the initial steps of steroid hormone synthesis. J Steroid Biochem Mol Biol. 2017;165:18–37. 7. Miller WL, Auchus RJ. The molecular biology, biochemistry, and physiology of human steroidogenesis and its disorders. Endocr Rev. 2011;32(1):81–151. 8. Conley AJ, Corbin CJ, Thomas JL, et al. Costs and consequences of cellular compartmentalization and substrate competition among human enzymes involved in androgen and estrogen synthesis. Biol Reprod. 2012;86(1):1–8. 9. Rainey WE, Carr BR, Sasano H, et al. Dissecting human adrenal androgen production. Trends Endocrinol Metab. 2002;13:234–239. 10. Kraemer FB. Adrenal cholesterol utilization. Mol Cell Endocrinol. 2007;265–266:42–45. 11. Sandhu J, Li S, Fairall L, et al. Aster proteins facilitate nonvesicular plasma membrane to ER cholesterol transport in mammalian cells. Cell. 2018;175(2):514–529. 12. Maxfield FR, Iaea DB, Pipalla NH. Role of STARD4 and NPC1 on intracellular sterol transport. Biochem Cell Biol. 2016;94:499–506. 13. Li NC, Fan J, Papadopoulos V. Sterol carrier protein-­2, a nonspecific lipid-­transfer protein, in intracellular cholesterol trafficking in testicular Leydig cells. PLoS One. 2016;11(2):e0149728. 14. Wilhelm LP, Tomasetto C, Alpy F. Touché! STARD3 and STARD3NL tether the ER to endosomes. Biochem Soc Trans. 2016;44(2):493–498. 15. Tugaeva KV, Sluchanko NN. Steroidogenic acute regulatory protein: structure, functioning, and regulation. Biochemistry (Mosc). 2019;84(suppl 1):S233–S253. 16. Hoekstra M, Van Eck M, Korporaal SJ. Genetic studies in mice and humans reveal new physiological roles for the high-­density lipoprotein receptor scavenger receptor class B type I. Curr Opin Lipidol. 2012;23:127–132. 17. Grøndahl C. Oocyte maturation. Basic and clinical aspects of in vitro maturation (IVM) with special emphasis of the role of FF-­ MAS. Dan Med Bull. 2008;55(1):1–16. 18. Keber R, Rozman D, Horvat S. Sterols in spermatogenesis and sperm maturation. J Lipid Res. 2013;54(1):20–33. 19. Keber R, Acimovic J, Majdic G, et al. Male germ cell-­ specific knockout of cholesterogenic cytochrome P450 lanosterol 14α-­ demethylase (Cyp51). J Lipid Res. 2013;54(6):1653–1661. 20. Illingworth DR, Corbin DK, Kemp ED, et al. Hormone changes during the menstrual cycle in abetalipoproteinemia: reduced luteal phase progesterone in a patient with homozygous hypobetalipoproteinemia. Proc Natl Acad Sci U S A. 1982;79:6685. 21. Parker Jr C, Illingworth DR, Bissonnette J, et al. Endocrine. changes during pregnancy in a patient with homozygous familial hypobetalipoproteinemia. N Engl J Med. 1986;314:557–560. 22. Plotkin D, Miller S, Nakajima S, et al. Lowering low density lipoprotein cholesterol with simvastatin, a hydroxy-­3-­methylglutaryl-­ coenzyme A reductase inhibitor, does not affect luteal function in premenopausal women. J Clin Endocrinol Metab. 2002;87:3155. 23. Laue L, Hoeg J, Barnes K, et al. The effect of mevinolin on steroidogenesis in patients with defects in the low density lipoprotein receptor pathway. J Clin Endocrinol Metab. 1987;64:531–535. 24. Porter FD, Herman GE. Malformation syndromes caused by disorders of cholesterol synthesis. J Lipid Res. 2011;52(1):6–34. 25. Andersson HC, Frentz J, Martinez JE, et al. Adrenal insufficiency in Smith-­ Lemli-­ Opitz syndrome. Am J Med Genet. 1999;82:382–384. 26. Shackleton CH produced by a pregnant woman carrying a SmithOpitz syn drome fetus. J Clin Endocrinol Metab. 1995;84:1157–1159.

27. Chang TY, Chang CCY, Chen D. Acyl-­coenzyme A: cholesterol acyltransferase. Annu Rev Biochem. 1997;66:613. 28. El-­Maouche D, Merke DP, Vogiatzi MG, et al. A phase 2, multicenter tudy of nevanimibe for the treatment of congenital adrenal hyperplasia. J Clin Endocrinol Metab. 2020;105(8):2771–2778. 29. Brasaemle DL. The perilipin family of structural lipid droplet proteins: stabilization of lipid droplets and control of lipolysis. J Lipid Res. 2007;48:2547. 30. Wang H, Sreenevasan U, Hu H, et al. Perilipin 5, a lipid droplet-­ associated protein, provides physical and metabolic linkage to mitochondria. J Lipid Res. 2011;52(12):2159–2168. 31. Ohta K, Sekiya M, Uozaki H, et al. Abrogation of neutral cholesterol ester hydrolytic activity causes adrenal enlargement. Biochem Biophys Res Commun. 2011;404(1):254–260. 32. Kraemer F, Shen W, Natu V, et al. Adrenal neutral cholesteryl ester hydrolase: identification, subcellular distribution, and sex difference. Endocrinology. 2002;143:801–806. 33. Li H, Brochu M, Wang SP, et al. Hormone-­sensitive lipase deficiency in mice causes lipid storage in the adrenal cortex and impaired corticosterone response to corticotropin stimulation. Endocrinology. 2002;143:3333–3340. 34.  Strauss III JF, Schuler LA, Rosenblum MF, et al. Cholesterol metabolism by ovarian tissue. Adv Lipid Res. 1981;18:99–157. 35. Andersen P, Zhang J, Su H, et al. Aster-­B coordinates with Arf1 to regulate mitochondrial cholesterol transport. Mol Metab. 2020;42:101055. 36. Ye J, DeBose-­Boyd RA. Regulation of cholesterol and fatty acid synthesis. Cold Spring Harb Perspect Biol. 2011;3(7):a004754. 37. Brown AJ, Jessup W. Oxysterols: sources, cellular storage and metabolism, and new insights into their roles in cholesterol homeostasis. Mol Aspects Med. 2009;30(3):111–122. 38. Cummins CL, Volle DH, Zhang Y, et al. Liver X receptors regulate adrenal cholesterol balance. J Clin Invest. 2006;116(7):1902–1912. 39. Tavori H, Rashid S, Fazio S. On the function and homeostasis of PCSK9: reciprocal interaction with LDLR and additional lipid effects. Atherosclerosis. 2015;238(2):264–270. 40. Devoto L, Fuentes A, Kohen P, et al. The human corpus luteum: life cycle and function in natural cycles. Fertil Steril. 2009;92(3):1067–1079. 41. Bogan RL, Hennebold JD. The reverse cholesterol transport system as a potential mediator of luteolysis in the primate corpus luteum. Reproduction. 2010;139(1):163–176. 42. Penning TM. Human hydroxysteroid dehydrogenases and pre-­ receptor regulation: insights into inhibitor design and evaluation. J Steroid Biochem Mol Biol. 2011;125(1–2):46–56. 43. Bose HS, Sugawara T, Strauss III JF, et al. International Congenital Lipoid Adrenal Hyperplasia Consortium: the pathophysiology and genetics of congenital lipoid adrenal hyperplasia. N Engl J Med. 1996;335:1870–1878. 44. Heyl BL, Tyrell DJ, Lambeth JD. Cytochrome P-­450scc-­substrate interactions: roles of the 3-­band side-­chain hydroxyls in binding to oxidized and reduced forms of the enzyme. J Biol Chem. 1986;261:2743. 45. Lambeth JD, Seybert DW, Kamin H. Phospholipid vesicle-­ reconstituted cytochrome P-­450scc: mutually facilitated binding of cholesterol and adrenodoxin. J Biol Chem. 1980;255:138. 46. Katsumata N, Ohtake M, Hojo T, et al. Compound heterozygous mutations in the cholesterol side-­ chain cleavage enzyme gene (CYP11A) cause congenital adrenal insufficiency in humans. J Clin Endocrinol Metab. 2002;87:3808–3813. 47. Hiort O, Holterhus PM, Werner R, et al. Homozygous disruption of P450 side-­chain cleavage (CYP11A1) is associated with prematurity, complete 46, XY sex reversal, and severe adrenal failure. J Clin Endocrinol Metab. 2005;90:538–541. 48. Hauffa B, Hiort O. P450 side-­chain cleavage deficiency—a rare cause of congenital adrenal hyperplasia. Endocr Dev. 2011;20:54–62. 49. Burkhard FZ, Parween S, Udhane SS, et al. P450 Oxidoreductase deficiency: analysis of mutations and polymorphisms. J Steroid Biochem Mol Biol. 2017;165:38–50. 50. Miller WL. The syndrome of 17,20 lyase deficiency. J Clin Endocrinol Metab. 2012;97(1):59–67. 51. Bhatt MR, Khatri Y, Rodgers RJ, et al. Role of cytochrome b5 in the modulation of the enzymatic activities of cytochrome P450 17α-­ hydroxylase/17,20-­lyase (P450 17A1). J Steroid Biochem Mol Biol. 2017;170:2–18. 52. Zhang L, Rodriguez H, Ohno S, et al. Serine phosphorylation of human P450c17 increases 17,20 lyase activity: implications for adrenarche and for polycystic ovary syndrome. Proc Natl Acad Sci U S A. 1995;92:10619–10623. 109.e1

109.e2 PART I  The Fundamentals of Reproduction 53. Cascio C, Prasad VV, Lin YY, et al. Detection of P450c17-­ independent pathways for dehydroepiandrosterone (DHEA) biosynthesis in brain glial tumor cells. Proc Natl Acad Sci U S A. 1998;95:2862–2867. 54. Sohl CD, Guengerich FP. Kinetic analysis of the three-­step steroid aromatase reaction of human cytochrome P450 19A1. J Biol Chem. 2010;285(23):17734–17743. 55. Kamat A, Hinshelwood MM, Murry BA, et al. Mechanisms in tissue-­specific regulation of estrogen biosynthesis in humans. Trends Endocrinol Metab. 2002;13:122–128. 56. Bulun SE. Clinical review 78: aromatase deficiency in women and men. Would you have predicted the phenotypes? J Clin Endocrinol Metab. 1996;81:867. 57. Belgorosky A, Guercio G, Pepe C, et al. Genetic and clinical spectrum of aromatase deficiency in infancy, childhood and adolescence. Horm Res. 2009;72(6):321–330. 58. Rochira V, Carani C. Aromatase deficiency in men: a clinical perspective. Nat Rev Endocrinol. 2009;5(10):559–568. 59. Simpson E, Clyne C, Rubin C. Aromatase: a brief overview. Annu Rev Physiol. 2002;64:93. 60. Stratakis CA, Vottero A, Brodie A, et al. The aromatase excess syndrome is associated with feminization of both sexes and autosomal dominant transmission of aberrant P450 aromatase gene transcription. J Clin Endocrinol Metab. 1998;83:1348–1357. 61. Bulun SE, Noble LS, Takayama K, et al. Endocrine. disorders associated with inappropriately high aromatase expression. J Steroid Biochem Mol Biol. 1997;61:133–139. 62. Shozu M, Sebastian S, Takayama K, et al. Estrogen excess associated with novel gain-­of-­function mutations affecting the aromatase gene. N Engl J Med. 2003;348:1855–1865. 63. Fukami M, Shozu M, Soneda S, et al. Aromatase excess syndrome: identification of cryptic duplications and deletions leading to gain of function of CYP19A1 and assessment of phenotypic determinants. J Clin Endocrinol Metab. 2011;96(6):E1035–E1043. 64. Imamichi Y, Yuhki KI, Orisaka M, et al. 11-­ketotestosterone is a major androgen produced in human gonads. J Clin Endocrinol Metab. 2016;101:3582–3591. 65. Krone N, Arlt W. Genetics of congenital adrenal hyperplasia. Best Pract Res Clin Endocrinol Metab. 2009;23(2):181–192. 66. White P, Rainey W. Steroid 11β-­hydroxylase isozymes. In: Mason JI, ed. Genetics of Steroid Biosynthesis and Function. Taylor & Francis; 2002:179–208. 67. Wendell A. Molecular genetics of 21-­hydroxylase deficiency. Endocr Dev. 2011;20:80–87. 68. Koppens PF, Hoogenboezem T, Degenhart HJ. Carriership of a defective tenascin-­ X gene in steroid 21-­ hydroxylase deficiency patients: TNXB-­TNXA hybrids in apparent large-­scale gene conversions. Hum Mol Genet. 2002;11:2581. 69. Nimkarn S, Lin-­ Su K, New MI. Steroid 21 hydroxylase deficiency congenital adrenal hyperplasia. Pediatr Clin North Am. 2011;58(5):1281–1300. 70. Saloniemi T, Jokela H, Strauss L, et al. The diversity of sex steroid action: novel functions of hydroxysteroid (17β) dehydrogenases as revealed by genetically modified mouse models. J Endocrinol. 2012:27–40. 71. Penning TM. The aldo-­keto reductases (AKRs): overview. Chem Biol Interact. 2015;234:236–246. 72. Lavery GG, Walker EA, Tiganescu A, et al. Steroid biomarkers and genetic studies reveal inactivating mutations in hexose-­6-­phosphate dehydrogenase in patients with cortisone reductase deficiency. J Clin Endocrinol Metab. 2008;93(10):3827–3832. 73. Seckl JR. 11β-­hydroxysteroid dehydrogenases: changing glucocorticoid action. Curr Opin Pharmacol. 2004;4(6):597–602. 74. Simard J, Ricketts M-­L, Gingras S, et al. Molecular biology of the 3β-­hydroxysteroid dehydrogenase/Δ5-­Δ4 isomerase gene family. Endocr Rev. 2005;26(4):525–582. 75. Lawson AJ, Walker EA, Lavery GG, et al. Cortisone-­reductase deficiency associated with heterozygous mutations in 11β-­ hydroxysteroid dehydrogenase type 1. Proc Natl Acad Sci U S A. 2011;108(10):4111–4116. 76. Kratschmar DV, Vuorinen A, Da Cunha T, et al. Characterization of activity and binding mode of glycyrrhetinic acid derivatives inhibiting characterization of dehydrogenase type 2. J Steroid Biochem Mol Biol. 2011;125(1–2):129–142.

77. Tanahashi T, Mune T, Morita H, et al. Glycyrrhizic acid suppresses type 2 11β-­ hydroxysteroid dehydrogenase expression in vivo. J Steroid Biochem Mol Biol. 2002;80:441–447. 78. Moeller G, Adamski J. Multifunctionality of human 17β-­ hydroxysteroid dehydrogenases. Mol Cell Endocrinol. 2006;248:47. 79. Lukacik P, Kavanagh KL, Oppermann U. Structure and function of human 17β-­hydroxysteroid dehydrogenases. Mol Cell Endocrinol. 2006;248:61. 80. Marchais-­Oberwinkler S, Henn C, Möller G, et al. 17β-­ Hydroxysteroid dehydrogenases (17β-­HSDs) as therapeutic targets: protein structures, functions, and recent progress in inhibitor development. J Steroid Biochem Mol Biol. 2011;125(1–2):66–82. 81. Hong Y, Chen S. Aromatase, estrone sulfatase, and 17β-­ hydroxysteroid dehydrogenase: structure-­ function studies and inhibitor development. Mol Cell Endocrinol. 2011;340(2):120–126. 82. Mendonca BB, Gomes NL, Costa EM, et al. 46,XY disorder of sex development (DSD) due to 17β-­hydroxysteroid dehydrogenase type 3 deficiency. J Steroid Biochem Mol Biol. 2017;165:79–85. 83. George MM, New MI, Ten S, et al. The clinical and molecular heterogeneity of 17βHSD-­3 enzyme deficiency. Horm Res Paediatr. 2010;74(4):229–240. 84. Luu-­The V, Dufort I, Pelletier G, et al. Type 5 17β-­hydroxysteroid dehydrogenase: its role in the formation of androgens in women. Mol Cell Endocrinol. 2001;171:77–82. 85. Nelson VL, Qin KN, Rosenfield RL, et al. The biochemical basis for increased testosterone production in theca cells propagated from patients with polycystic ovary syndrome. J Clin Endocrinol Metab. 2001;86:5925–5933. 86. Flück CE, Meyer-­Böni M, Pandey AV, et al. Why boys will be boys: two pathways of fetal testicular androgen biosynthesis are needed for male sexual differentiation. Am J Hum Genet. 2011;89(2):201–218. 87. Russell DW, Wilson JD. Steroid 5α-­ reductase: two genes/two enzymes. Annu Rev Biochem. 1994;63:25. 88. Wilson JD, Griffin JE, Russell DW. Steroid 5-­reductase 2 deficiency. Endocr Rev. 1993;14:577. 89. Mendonca BB, Batista RL, Domenice S, et al. Steroid 5α-­reductase 2 deficiency. J Steroid Biochem Mol Biol. 2016;163:206–211. 90. Mahendroo MS, Cala K, Russell DW. 5α-­Reduced androgens play a key role in murine parturition. Mol Endocrinol. 1996;10:380. 91. Mahendroo MS, Cala KM, Landrum DP, et al. Fetal death in mice lacking 5α-­reductase type 1 caused by estrogen excess. Mol Endocrinol. 1997;11:917–927. 92. Kondo K, Kai M, Setoguchi Y, et al. Cloning and expression of cDNA of human 4-­4-­3-­oxosteroid-­5β-­reductase and substrate specificity of the expressed enzyme. Eur J Biochem. 1994;219:357. 93. Negishi M, Pedersen LG, Petrotchenko E, et al. Structure and function of sulfotransferases. Arch Biochem Biophys. 2001;390:149–157. 94. Fuda H, Lee YC, Shimizu C, et al. Mutational analysis of human hydroxysteroid sulfotransferase SULT2B1 isoforms reveals that exon 1B of the SULT2B1 gene produces cholesterol sulfotransferase, whereas exon 1A yields pregnenolone sulfotransferase. J Biol Chem. 2002;277:36161. 95. Suzuki T, Miki Y, Nakamura Y, et al. Steroid sulfatase and estrogen sulfotransferase in human carcinomas. Mol Cell Endocrinol. 2011;340(2):148–153. 96. Qian YM, Sun XJ, Tong MH, et al. Targeted disruption of the mouse estrogen sulfotransferase gene reveals a role of estrogen metabolism in intracrine and paracrine estrogen regulation. Endocrinology. 2001;142:5342–5350. 97. Kester MH, Bulduk S, Tibboel D, et al. Potent inhibition of estrogen sulfotransferase by hydroxylated PCB metabolites: a novel pathway explaining the estrogenic activity of PCBs. Endocrinology. 2000;141:1897–1900. 98. Purohit A, Woo LW, Potter BV. Steroid sulfatase: a pivotal player in estrogen synthesis and metabolism. Mol Cell Endocrinol. 2011;340(2):154–160. 99. Yen PH, Li XM, Tsai SP, et al. Frequent deletions of the human X chromosome distal short arm result from recombination between low copy repetitive elements. Cell. 1990;61:603–610. 100. Ballabio A, Carrozzo R, Parenti G, et al. Molecular heterogeneity of steroid sulfatase deficiency: a multicenter study on 57 unrelated patients, at DNA and protein levels. Genomics. 1989;4:36–40. 101. Foster PA, Purohit A. Steroid sulfatase inhibitors for estrogen-­and androgen-­dependent cancers. J Endocrinol. 2012;212(2):99–110.

CHAPTER 4  Steroid Hormones and Other Lipid Molecules Involved in Human Reproduction 102. Albert C, Barbier O, Vallee M, et al. Distribution of uridine diphosphate-­glucuronosyltransferase (UGT) expression and activity in cynomolgus monkey tissues: evidence for differential expression of steroid-­conjugating UGT enzymes in steroid target tissues. Endocrinology. 2000;141:2472–2480. 103. Turgeon D, Carrier JS, Levesque E, et al. Relative enzymatic activity, protein stability, and tissue distribution of human steroid-­metabolizing UGT2B subfamily members. Endocrinology. 2001;142:778–787. 104. Levesque E, Turgeon D, Carrier JS, et al. Isolation and characterization of the UGT2B28 cDNA encoding a novel human steroid conjugating UDP-­ glucuronosyltransferase. Biochemistry. 2001;40:3869–3881. 105. Kohalmy K, Vrzal R. Regulation of phase II biotransformation enzymes by steroid hormones. Curr Drug Metab. 2011;12(2):104–123. 106. Barbier O, Bélanger A. Inactivation of androgens by UDP-­ glucuronosyltransferases in the human prostate. Best Pract Res Clin Endocrinol Metab. 2008;22(2):259–270. 107. Auchus RJ. The backdoor pathway to dihydrotestosterone. Trends Endocrinol Metab. 2004;15(9):432–438. 108. Turcu AF, Rege J, Auchus RJ, Rainey WE. 11-­Oxygenated androgens in health and disease. Nat Rev Endocrinol. 2020;16(5):284–296. 109. Barnard L, Schiffer L, Louw du-­Toit R, et al. 11-­oxygenated estrogens are a novel class of human estrogens but do not contribute to the circulating estrogen pool. Endocrinology. 2021;162(3). 110. Weisz J, Clawson GA, Creveling CR. Biogenesis and inactivation of catecholestrogens. Adv Pharmacol. 1998;42:828. 111. Cavalieri E, Frenkel K, Liehr JG, et al. Estrogens as endogenous genotoxic agents: DNA adducts and mutations. J Natl Cancer Inst Monogr. 2000;27:75–93. 112. Mooberry SL. New insights into 2-­methoxyestradiol, a promising antiangiogenic and antitumor agent. Curr Opin Oncol. 2003;15:425. 113. Kanasaki K, Palmsten K, Sugimoto H, et al. Deficiency in catechol-­ O-­ methyltransferase and 2-­ methoxyoestradiol is associated with pre-­eclampsia. Nature. 2008;453(7198):1117–1121. 114. Perez-­Sepulveda A, España-­Perrot PP, Norwitz ER, et al. Metabolic pathways involved in 2-­methoxyestradiol synthesis and their role in preeclampsia. Reprod Sci. 2013;20(9):1020–1029. 115. Henríquez S, Kohen P, Xu X, et al. Estrogen metabolites in human corpus luteum physiology: differential effects on angiogenic activity. Fertil Steril. 2016;106(1):230–237.e1. 116. Hochberg RB, Pahuja SL, Zielinski JE, et al. Steroidal fatty acid esters. J Steroid Biochem Mol Biol. 1991;40:577–585. 117. Vihma V, Tikkanen MJ. Fatty acid esters of steroids: synthesis and metabolism in lipoproteins and adipose tissue. J Steroid Biochem Mol Biol. 2011;124(3–5):65–76. 118. Chalbot S, Morfin R. Dehydroepiandrosterone metabolites and their interactions in humans. Drug Metabol Drug Interact. 2006;22(1):1–23. 119. Foster SJ, Marshall DE, Houghton E, et al. Investigations into the biosynthetic pathways for classical and ring B-­unsaturated oestrogens in equine placental preparations and allantochorionic tissues. J Steroid Biochem Mol Biol. 2002;82(4–5):401–411. 120. Nandi A, Sinha N, Ong E, et al. Is there a role for vitamin D in human reproduction? Horm Mol Biol Clin Investig. 2016;25(1):15–28. 121. Christakos S, Ajibade DV, Dhawan P, et al. Vitamin D: metabolism. Endocrinol Metab Clin North Am. 2010;39(2):243–253. 122. Henry HL. Regulation of vitamin D metabolism. Best Pract Res Clin Endocrinol Metab. 2011;25(4):531–541. 123. Miller WL. Genetic disorders of Vitamin D biosynthesis and degradation. J Steroid Biochem Mol Biol. 2017;165:101–108. 124. Schimmer BP, White PC. Minireview: steroidogenic factor 1: its roles in differentiation, development, and disease. Mol Endocrinol. 2010;24(7):1322–1337. 125. Ferraz-­de-­Souza B, Lin L, Achermann JC. Steroidogenic factor-­ 1 (SF-­ 1, NR5A1) and human disease. Mol Cell Endocrinol. 2011;336(1–2):198–205. 126. Fayard E, Auwerx J, Schoonjans K. LRH-­1: an orphan nuclear receptor involved in development, metabolism and steroidogenesis. Trends Cell Biol. 2004;14:250. 127. Krylova IN, Sablin EP, Moore J, et al. Structural analyses reveal phosphatidyl inositols as ligands for the NR5 orphan receptors SF-­1 and LRH-­1. Cell. 2005;120:343–355. 128. Martin LJ, Taniguchi H, Robert NM, et al. GATA factors and the nuclear receptors, steroidogenic factor 1/liver receptor homolog 1, are key mutual partners in the regulation of the human 3β-­hydroxysteroid dehydrogenase type 2 promoter.

109.e3

129. King SR, Lavoie HA. Gonadal transactivation of STARD1, CYP11A1 and HSD3B. Front Biosci. 2012;1(17):824–846. 130. Tuckey RC. Progesterone synthesis by the human placenta. Placenta. 2005;26:273. 131. Luu-­The V, Labrie F. The intracrine sex steroid biosynthesis pathways. Prog Brain Res. 2010;181:177–192. 132. Lathe R, Seckl J. Neurosteroids and brain sterols. In: Mason JI, ed. Genetics of Steroid Biosynthesis and Function. Taylor & Francis; 2002:407–474. 133. Attar E, Bulun SE. Aromatase and other steroidogenic genes in endometriosis: translational aspects. Hum Reprod Update. 2006;12:49–56. 134. Mellon SH, Griffin LD. Neurosteroids: biochemistry and clinical significance. Trends Endocrinol Metab. 2002;13:35. 135. Pelletier G. Steroidogenic enzymes in the brain: morphological aspects. Prog Brain Res. 2010;181:193–207. 136. Deplewski D, Rosenfield RL. Role of hormones in pilosebaceous unit development. Endocr Rev. 2002;21:363. 137. Andersson S. Steroidgenic enzymes in skin. Eur J Dermatol. 2001;11(4):293–295. 138. Couzinet B, Meduri G, Lecce MG, et al. The postmenopausal ovary is not a major androgen-­producing gland. J Clin Endocrinol Metab. 2001;86:5060. 139. Tait JF. The use of isotopic steroids for the measurement of production rates in vivo. J Clin Endocrinol Metab. 1963;23:1285. 140. Milewich L, Parker PS, MacDonald PC. Testosterone metabolism by human lung tissue. J Steroid Biochem. 1978;9:29. 141. Milewich L, Smith SL, MacDonald PC. Nonrespiratory functions of the human lung: in vitro metabolism of tritium-­labeled progesterone and pregnenolone. J Clin Endocrinol Metab. 1980;50:507. 142. Dunn JF, Nisula BC, Rodbard D. Transport of steroid hormones: binding of 21 endogenous steroids to both testosterone-­binding globulin and corticoid-­binding globulin in human plasma. J Clin Endocrinol Metab. 1981;53:58. 143. Sitteri PK, Murai JT, Hammond GL, et al. The serum transport of steroid hormones. Recent Prog Horm Res. 1982;38:457. 144. Manni A, Pardridge WM, Cefalu W, et al. Bioavailability of albumin-­ bound testosterone. J Clin Endocrinol Metab. 1985;61:705–710. 145. Baird DT, Horton R, Longcope C, et al. Steroid dynamics under steady-­state conditions. Recent Prog Horm Res. 1969;25:611–664. 146. Hogeveen KN, Cousin P, Pugeat M, et al. Human sex hormone-­ binding globulin variants associated with hyperandrogenism and ovarian dysfunction. J Clin Invest. 2002;109:973–981. 147. Ukkola O, Rankinen T, Gagnon J, et al. A genome-­wide linkage scan for steroids and SHBG levels in black and white families: the HERITAGE Family Study. J Clin Endocrinol Metab. 2002;87:3708–3720. 148. Cousin P, Calemard-­Michel L, Lejeune H, et al. Influence of SHBG gene pentanucleotide TAAAA repeat and D327N polymorphism on serum sex hormone-­binding globulin concentration in hirsute women. J Clin Endocrinol Metab. 2004;89:917–924. 149. Le TN, Nestler JE, Strauss 3rd JF, et al. Sex hormone-­binding globulin and type 2 diabetes mellitus. Trends Endocrinol Metab. 2012;23(1):32–40. 150. Hammond GL. Diverse roles for sex hormone-­binding globulin in reproduction. Biol Reprod. 2011;85(3):431–441. 151. Craig ZR, Wang W, Flaws JA. Endocrine-­disrupting chemicals in ovarian function: effects on steroidogenesis, metabolism and nuclear receptor signaling. Reproduction. 2011;142:633–646. 152. Shaw MA, Nicholls PJ, Smith HJ. Aminoglutethimide and ketoconazole: historical perspectives and future prospects. J Steroid Biochem. 1988;31:137. 153. Bryce A, Ryan CJ. Development and clinical utility of abiraterone acetate as an androgen synthesis inhibitor. Clin Pharmacol Ther. 2012;91(1):101–108. 154. Stein MN, Patel N, Bershadskiy A, et al. Androgen synthesis inhibitors in the treatment of castration-­resistant prostate cancer. Asian J Androl. 2014;16(3):387–400. 155. Engelhardt D, Weber MM. Therapy of Cushing’s syndrome with steroid biosynthesis inhibitors. J Steroid Biochem Mol Biol. 1994;49:261. 156. Chumsri S, Howes T, Bao T, et al. Aromatase, aromatase inhibitors, and breast cancer. J Steroid Biochem Mol Biol. 2011;125(1–2):13–22. 157. Bull HG, Gacia-­Calvo M, Andersson S, et al. Mechanism-­based inhibition of human steroid 5α-­reductase by finasteride: enzyme-­ catalyzed formation of NADP-­dihydrofinasteride, a potent bisub. 1996;118:2359.

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109.e4 PART I  The Fundamentals of Reproduction 158. Goldman AS. Further studies of steroidal inhibitors of delpha5, 3beta-­ hydroxysteroid dehydrogenase and delta5-­ delta4, 3-­ketosteroid isomerase in Pseudomonas testosteroni and in bovine adrenals. J Clin Endocrinol Metab. 1968;28:1539. 159. Christiansen RG, Neumann HG, Salvador UJ, et al. Steroidogenesis inhibitors: adrenal inhibitory and interceptive activity of trilostane and related compounds. J Med Chem. 1984;27:928–931. 160. Thomas MP, Potter BV. The structural biology of oestrogen metabolism. J Steroid Biochem Mol Biol. 2013;137:27–49. 161. de Gooyer ME, Kleyn GT, Smits KC, et al. Tibolone: a compound with tissue specific inhibitory effects on sulfatase. Mol Cell Endocrinol. 2001;183:55–62. 162. von Euler US. History and development of prostaglandins. Gen Pharmacol. 1983;14:3–6. 163. Funk CD. Prostaglandins and leukotrienes: advances in eicosanoid biology. Science. 2001;294:1871–1875. 164. Oates JA. The 1982 Nobel prize in physiology or medicine. Science. 1982;218:765–768. 165. Brash AR. Arachidonic acid as a bioactive molecule. J Clin Invest. 2001;107:1339–1345. 166. Fitzpatrick FA, Soberman R. Regulated formation of eicosanoids. J Clin Invest. 2001;107:1347–1351. 167. Burke JE, Dennis EA. Phospholipase A2 structure/function, mechanism, and signaling. J Lipid Res. 2009;50(suppl):S237–S242. 168. Murakami M, Kambe T, Shimbara S, et al. Functional coupling between various phospholipase A2s and cyclooxygenases in immediate and delayed prostanoid biosynthetic pathways. J Biol Chem. 1999;274:3103–3115. 169. Tanioka T, Nakatani Y, Semmyo N, et al. Molecular identification of cytosolic prostaglandin E2 synthase that is functionally coupled with cyclooxygenase-­1 in immediate prostaglandin E2 biosynthesis. J Biol Chem. 2000;275:32775–32782. 170. Murakami M, Naraba H, Tanioka T, et al. Regulation of prostaglandin E2 biosynthesis by inducible membrane-­associated prostaglandin E2 synthase that acts in concert with cyclooxygenase-­2. J Biol Chem. 2000;275:32783–32792. 171. Cho YY, Kang MJ, Sone H, et al. Abnormal uterus with polycysts, accumulation of uterine prostaglandins, and reduced fertility in mice heterozygous for acyl-­CoA synthetase 4 deficiency. Biochem Biophys Res Commun. 2001;284:993–997. 172. Smith WL, DeWitt DL, Garavito RM. Cyclooxygenases: structural, cellular, and molecular biology. Annu Rev Biochem. 2000;69:145–182. 173. Smith W. Molecular biology of prostanoid biosynthetic enzymes and receptors. Adv Exp Med Biol. 1997;400B:989–1011. 174. Brash AR. Lipoxygenases: occurrence, functions, catalysis, and acquisition of substrate. J Biol Chem. 1999;274:23679–23682. 175. Peters-­Golden M, Henderson Jr WR. Leukotrienes. N Engl J Med. 2007;357(18):1841–1854. 176. Serhan CN. Controlling the resolution of acute inflammation: a new genus of dual anti-­inflammatory and proresolving mediators. J Periodontol. 2008;79:1520–1526. 177. Spector AA. Arachidonic acid cytochrome P450 epoxygenase pathway. J Lipid Res. 2009;50:S52–S56. 178. Holla VR, Makita K, Zaphiropoulos PG, et al. The kidney cytochrome P-­450 2C23 arachidonic acid epoxygenase is upregulated during dietary salt loading. J Clin Invest. 1999;104:751–760. 179. Holla VR, Adas F, Imig JD, et al. Alterations in the regulation of androgen-­ sensitive Cyp 4a monooxygenases cause hypertension. Proc Natl Acad Sci U S A. 2001;98:5211–5216. 180. Catella F, Lawson JA, Fitzgerald DJ, et al. Endogenous biosynthesis of arachidonic acid epoxides in humans: increased formation in pregnancy-­induced hypertension. Proc Natl Acad Sci U S A. 1990;87:5893–5897. 181. Node K, Huo Y, Ruan X, et al. Anti-­ inflammatory properties of cytochrome P450 epoxygenase-­ derived eicosanoids. Science. 1999;285:1276–1279. 182. Rosen EM, van ‘t Erve TJ, Boss J, et al. Urinary oxidative stress biomarkers and accelerated time to spontaneous delivery. Free Radic Biol Med. 2019;130:419–425. 183. Coviello C, Tataranno ML, Corsini I, et al. Isoprostanes as biomarker for patent ductus arteriosus in preterm infants. Front Pediatr. 2020;8:555. 184. Signorini C, Moretti E, Collodel G. Role of isoprostanes in human male infertility. Syst Biol Reprod Med. 2020;66(5):291–299. 185. Roberts II LJ, Morrow JD. Isoprostanes: novel markers of endogenous lipid peroxidation and potential mediators of oxidant injury. Ann N Y Acad Sci. 1994;744:237–242.

186. Milne GL, Yin H, Hardy KD, et al. Isoprostane generation and function. Chem Rev. 2011;111(10):5973–5996. 187. Kunapuli P, Lawson JA, Rokach JA, et al. Prostaglandin F2alpha (PGF2alpha) and the isoprostane, 8, 12-­iso-­isoprostane F2alpha-­ III,induce cardiomyocyte hypertrophy: differential activation of downstream signaling pathways. J Biol Chem. 1998;273:22442–22452. 188. Davi G, Guagnano MT, Ciabattoni G, et al. Platelet activation in obese women: role of inflammation and oxidant stress. JAMA. 2002;288:2008–2014. 189. Meagher EA, Barry OP, Burke A, et al. Alcohol-­ induced generation of lipid peroxidation products in humans. J Clin Invest. 1999;104:805–813. 190. Smith WL, Urade Y, Jakobsson PJ. Enzymes of the cyclooxygenase pathways of prostanoid biosynthesis. Chem Rev. 2011;111(10):5821–5865. 191. Harris RC, McKanna JA, Akai Y, et al. Cyclooxygenase-­2 is associated with the macula densa of rat kidney and increases with salt restriction. J Clin Invest. 1994;94:2504–2510. 192. Funk CD, Radmark O, Fu JY, et al. Molecular cloning and amino acid sequence of leukotriene A4 hydrolase. Proc Natl Acad Sci U S A. 1987;84:6677–6681. 193. Samuelsson B, Morgenstern R, Jakobsson PJ. Membrane prostaglandin E synthase-­1: a novel therapeutic target. Pharmacol Rev. 2007;59(3):207–224. 194. Parent J, Fortier MA. Expression and contribution of three different isoforms of prostaglandin E synthase in the bovine endometrium. Biol Reprod. 2005;73:36–44. 195. Morrow JD, Awad JA, Boss HJ, et al. Non-­cyclooxygenase-­derived prostanoids (F2-­isoprostanes) are formed in situ on phospholipids. Proc Natl Acad Sci U S A. 1992;89:10721–10725. 196. Samuelsson B. Leukotrienes: mediators of immediate hypersensitivity reactions and inflammation. Science. 1983;220:568–575. 197. Haeggström JZ, Funk CD. Lipoxygenase and leukotriene pathways: biochemistry, biology, and roles in disease. Chem Rev. 2011;111(10):5866–5898. 198. Bäck M, Dahlén SE, Drazen JM, et al. International Union of Basic and Clinical Pharmacology. LXXXIV: leukotriene receptor nomenclature, distribution, and pathophysiological functions. Pharmacol Rev. 2011;63(3):539–584. 199. Yerushalmi GM, Markman S, Yung Y, et al. The prostaglandin transporter (PGT) as a potential mediator of ovulation. Sci Transl Med. 2016;8(338):338ra68. 200. Peters-­Golden M, McNish RW. Redistribution of 5-­lipoxygenase and cytosolic phospholipase A2 to the nuclear fraction upon macrophage activation. Biochem Biophys Res Commun. 1993;196:147–153. 201. Chen XS, Funk CD. The N-­ terminal “beta-­ barrel” domain of 5-­lipoxygenase is essential for nuclear membrane translocation. J Biol Chem. 2001;276:811–818. 202. Evans JF, Ferguson AD, Mosley RT, et al. What’s all the FLAP about?: 5-­lipoxygenase-­activating protein inhibitors for inflammatory diseases. Trends Pharmacol Sci. 2008;29(2):72–78. 203. Roizen JD, Asada M, Tong M, Tai HH, Muglia LJ. Early pregnancy loss in 15-­ hydroxyprostaglandin dehydrogenase knockout (15-­HPGD-­/-­) mice due to requirement for embryo 15-­HPGD activity. Sci Rep. 2019;9(1):17612. 204. Penrose JF, Austen KF. The biochemical, molecular, and genomic aspects of leukotriene C4 synthase. Proc Assoc Am Physicians. 1999;111:537–546. 205. Folco G, Murphy RC. Eicosanoid transcellular biosynthesis: from cell-­ cell interactions to in vivo tissue responses. Pharmacol Rev. 2006;58:375–388. 206. Schuster VL. Molecular mechanisms of prostaglandin transport. Annu Rev Physiol. 1998;60:221–242. 207. Reid G, Wielinga P, Zelcer N, et al. The human multidrug resistance protein MRP4 functions as a prostaglandin efflux transporter and is inhibited by nonsteroidal antiinflammatory drugs. Proc Natl Acad Sci U S A. 2003;100:9244–9249. 208. Cole SP, Deeley RG. Transport of glutathione and glutathione conjugates by MRP1. Trends Pharmacol Sci. 2006;27:438–446. 209. Ensor CM, Yang JY, Okita RT, et al. Cloning and sequence analysis of the cDNA for human placental NAD(+)-­dependent 15-­hydroxyprostaglandin dehydrogenase. J Biol Chem. 1990;265:14888–14891. 210. FitzGerald GA, Loll P. COX in a crystal ball: current status and future promise of prostaglandin research. J Clin Invest.

CHAPTER 4  Steroid Hormones and Other Lipid Molecules Involved in Human Reproduction 211. Yu Y, Fan J, Hui Y, et al. Targeted cyclooxygenase gene (ptgs) exchange reveals discriminant isoform functionality. J Biol Chem. 2007;282(2):1498–1506. 212. Chen W, Pawelek TR, Kulmacz RJ. Hydroperoxide dependence and cooperative cyclooxygenase kinetics in prostaglandin H synthase-­1 and -­2. J Biol Chem. 1999;274:20301–20306. 213. FitzGerald GA, Patrono C. The coxibs, selective inhibitors of cyclooxygenase-­2. N Engl J Med. 2001;345:433–442. 214. Grosser T, Ricciotti E, FitzGerald GA. The cardiovascular pharmacology of nonsteroidal anti-­inflammatory drugs. Trends Pharmacol Sci. 2017;38(8):733–748. 215. Grosser T, Yusuff S, Cheskis E, et al. Developmental expression of functional cyclooxygenases in zebrafish. Proc Natl Acad Sci U S A. 2002;99:8418–8423. 216. Langenbach R, Morham SG, Tiano HF, et al. Prostaglandin synthase 1 gene disruption in mice reduces arachidonic acid-­induced inflammation and indomethacin-­ induced gastric ulceration. Cell. 1995;83:483–492. 217. Morham SG, Langenbach R, Loftin CD, et al. Prostaglandin synthase 2 gene disruption causes severe renal pathology in the mouse. Cell. 1995;83:473–482. 218. Dinchuk JE, Car BD, Focht RJ, et al. Renal abnormalities and an altered inflammatory response in mice lacking cyclooxygenase II. Nature. 1995;378:406–409. 219. Lim H, Paria BC, Das SK, et al. Multiple female reproductive failures in cyclooxygenase 2-­deficient mice. Cell. 1997;91:197–208. 220. McAdam BF, Catella-­Lawson F, Mardini IA, et al. Systemic biosynthesis of prostacyclin by cyclooxygenase (COX)-­2: the human pharmacology of a selective inhibitor of COX-­2. Proc Natl Acad Sci U S A. 1999;96:272–277. 221. Woodward DF, Jones RL, Narumiya S. International Union of Basic and Clinical Pharmacology. LXXXIII: classification of prostanoid receptors, updating 15 years of progress. Pharmacol Rev. 2011;63(3):471–538. 222. Narumiya S, Furuyashiki T. Fever, inflammation, pain and beyond: prostanoid receptor research during these 25 years. FASEB J. 2011;25(3):813–818. 223. Coleman RA, Smith WL, Narumiya S. International Union of Pharmacology classification of prostanoid receptors: properties, distribution, and structure of the receptors and their subtypes. Pharmacol Rev. 1994;46:205–229. 224. Hirata M, Hayashi Y, Ushikubi F, et al. Cloning and expression of cDNA for a human thromboxane A2 receptor. Nature. 1991;349:617–620. 225. Narumiya S. Molecular diversity of prostanoid receptors: subtypes and isoforms of prostaglandin E receptor. Adv Exp Med Biol. 1997:207–213. 226. Pierce KL, Bailey TJ, Hoyer PB, et al. Cloning of a carboxyl-­ terminal isoform of the prostanoid FP receptor. J Biol Chem. 1997;272:883–887. 227. Raychowdhury MK, Yukawa M, Collins LJ, et al. Alternative splicing produces a divergent cytoplasmic tail in the human endothelial thromboxane A2 receptor. J Biol Chem. 1994;269:19256–19261. 228. Sugimoto Y, Inazumi T, Tsuchiya S. Roles of prostaglandin receptors in female reproduction. J Biochem. 2015;157(2):73–80. 229. Kanaoka Y, Maekawa A, Austen KF. Identification of GPR99 protein as a potential third cysteinyl leukotriene receptor with a preference for leukotriene E4 ligand. J Biol Chem. 2013;288(16):10967–10972. 230. Hirai H, Tanaka K, Yoshie O, et al. Prostaglandin D2 selectively induces chemotaxis in T helper type 2 cells, eosinophils, and basophils via seven-­ transmembrane receptor CRTH2. J Exp Med. 2001;193:255–261. 231. Monneret G, Gravel S, Diamond M, et al. Prostaglandin D2 is a potent chemoattractant for human eosinophils that acts via a novel DP receptor. Blood. 2001;98:1942–1948. 232. Monneret G, Li H, Vasilescu J, et al. 15-­ Deoxy-­ delta 12, 14-­prostaglandins D2 and J2 are potent activators of human eosinophils. J Immunol. 2002;168:3563–3569. 233. Yamamoto T, Miyata J, Arita M, Fukunaga K, Kawana A. Current state and future prospect of the therapeutic strategy targeting cysteinyl leukotriene metabolism in asthma. Respir Investig. 2019;57(6):534–543. 234. Kanaoka Y, Austen KF. Roles of cysteinyl leukotrienes and their receptors in immune cell-­ related functions. Adv Immunol. 2019;142:65–84. 235. Nakamura M . Leukotriene receptors. 2011;111(10):6231–6298.

109.e5

236. Chandrasekharan JA, Sharma-­Walia N. Lipoxins: nature’s way to resolve inflammation. J Inflamm Res. 2015;8:181–189. 237. Lim H, Dey SK. Minireview: a novel pathway of prostacyclin signaling-­ hanging out with nuclear receptors. Endocrinology. 2002;143:3207–3210. 238. Yokomizo T, Kato K, Terawaki K, et al. A second leukotriene B(4) receptor, BLT2: a new therapeutic target in inflammation and immunological disorders. J Exp Med. 2000;192:421–432. 239. Labat C, Ortiz JL, Norel X, et al. A second cysteinyl leukotriene receptor in human lung. J Pharmacol Exp Ther. 1992;263:800–805. 240. Lynch KR, O’Neill GP, Liu Q, et al. Characterization of the human cysteinyl leukotriene CysLT1 receptor. Nature. 1999;399:789–793. 241. Heise CE, O’Dowd BF, Figueroa DJ, et al. Characterization of the human cysteinyl leukotriene 2 receptor. J Biol Chem. 2000;275:30531–30536. 242. Barajas-­Espinosa A, Ni NC, Yan D, et al. The cysteinyl leukotriene 2 receptor mediates retinal edema and pathological neovascularization in a murine model of oxygen-­induced retinopathy. FASEB J. 2012;26(3):1100–1109. 243. Yu K, Bayona W, Kallen CB, et al. Differential activation of peroxisome proliferator-­activated receptors by eicosanoids. J Biol Chem. 1995;270:23975–23983. 244. Forman BM, Tontonoz P, Chen J, et al. 15-­ Deoxy-­ delta 12, 14-­prostaglandin J2 is a ligand for the adipocyte determination factor PPAR gamma. Cell. 1995;83:803–812. 245. Kliewer SA, Lenhard JM, Willson TM, et al. A prostaglandin J2 metabolite binds peroxisome proliferator-­activated receptor gamma and promotes adipocyte differentiation. Cell. 1995;83:813–819. 246. Devchand PR, Keller H, Peters JM, et al. The PPARalpha-­ leukotrieneB4 pathway to inflammation control. Nature. 1996;384:39–43. 247. Gupta RA, Tan J, Krause WF, et al. Prostacyclin-­mediated activation of peroxisome proliferator-­activated receptor delta in colorectal cancer. Proc Natl Acad Sci U S A. 2000;97:13275–13280. 248. Hatae T, Wada M, Yokoyama C, et al. Prostacyclin-­dependent apoptosis mediated by PPAR delta. J Biol Chem. 2001;276:46260–46267. 249. Lim H, Gupta RA, Ma WG, et al. Cyclo-­oxygenase-­2-­derived prostacyclin mediates embryo implantation in the mouse via PPARdelta. Genes Dev. 1999;13:1561–1574. 250. FIGO working group on prevention of Unsafe abortion and its consequences; International Federation of gynecology and Obstetrics: the combination of mifepristone and misoprostol for the termination of pregnancy. Int J Gynaecol Obstet. 2011;115(1):1–4. 251. Chen QJ, Hou SP, Meads C, et al. Mifepristone in combination with prostaglandins for termination of 10-­16 weeks’ gestation: a systematic review. Eur J Obstet Gynecol Reprod Biol. 2011;159(2):247–254. 252. Taher SE, Inder JW, Soltan SA, et al. Prostaglandin E2 vaginal gel or tablets for the induction of labour at term: a randomised controlled trial. BJOG. 2011;118:719–725. 253. Murdoch WJ, Hansen TR, McPherson LA. A review: role of eicosanoids in vertebrate ovulation. Prostaglandins. 1993;46:85–115. 254. McCracken JA, Carlson JC, Glew ME, et al. Prostaglandin F2 identified as a luteolytic hormone in sheep. Nat New Biol. 1972;238:129–134. 255. Moeljono MP, Bazer FW, Thatcher WW. A study of prostaglandin F2alpha as the luteolysin in swine: I. Effect of prostaglandin F2alpha in hysterectomized gilts. Prostaglandins. 1976;11:737–743. 256. Heymann MA, Rudolph AM, Silverman NH. Closure of the ductus arteriosus in premature infants by inhibition of prostaglandin synthesis. N Engl J Med. 1976;295:530–533. 257. Mielke G, Gonser M. Closure of the human fetal ductus arteriosus. Am J Obstet Gynecol. 1997;176:495–496. 258. Hedin L, Gaddy-­Kurten D, Kurten R, et al. Prostaglandin endoperoxide synthase in rat ovarian follicles: content, cellular distribution, and evidence for hormonal induction preceding ovulation. Endocrinology. 1987;121:722–731. 259. Wong WY, DeWitt DL, Smith WL, et al. Rapid induction of prostaglandin endoperoxide synthase in rat preovulatory follicles by luteinizing hormone and cAMP is blocked by inhibitors of transcription and translation. Mol Endocrinol. 1989;3:1714–1723. 260. Wong WY, Richards JS. Evidence for two antigenically distinct molecular weight variants of prostaglandin H synthase in the rat ovary. Mol Endocrinol. 1991;5:1269–1279. 261. Colley R, Wilson J. How to Begin hyperalimentation therapy: meeting patients’ nutritional needs with hyperalimentation. Nursing. 1979;9(5):76–83. 262. Sirois J, Levy LO, Simmons DL, et al. Characterization and hormonal regulation of the promoter of the rat prostaglandin endoperoxide synthase 2 gene in granulosa cells: identification of functional and protein-­binding regions. J Biol Chem. 1993;268:12199–12206.

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109.e6 PART I  The Fundamentals of Reproduction 263. Sirois J, Dore M. The late induction of prostaglandin G/H synthase­2 in equine preovulatory follicles supports its role as a determinant of the ovulatory process. Endocrinology. 1997;138:4427–4434. 264. Duffy DM. Novel contraceptive targets to inhibit ovulation: the prostaglandin E2 pathway. Hum Reprod Update. 2015;21(5):652–670. 265. Duffy DM, Stouffer RL. Follicular administration of a cyclooxygenase inhibitor can prevent oocyte release without alteration of normal luteal function in rhesus monkeys. Hum Reprod. 2002;17:2825–2831. 266. Richards JS. Sounding the alarm: does induction of prostaglandin endoperoxide synthase-­2 control the mammalian ovulatory clock? Endocrinology. 1997;138:4047–4048. 267. Richards JS, Russell DL, Ochsner S, et al. Ovulation: new dimensions and new regulators of the inflammatory-­like response. Annu Rev Physiol. 2002;64:69–92. 268. Gaytán F, Tarradas E, Bellido C, Morales C, Sánchez-­Criado JE. Prostaglandin E(1) inhibits abnormal follicle rupture and restores ovulation in indomethacin-­ treated rats. Biol Reprod. 2002;67(4):1140–1147. 269. Kim SO, Harris SM, Duffy DM. Prostaglandin E2 (EP) receptors mediate PGE2-­specific events in ovulation and luteinization within primate ovarian follicles. Endocrinology. 2014;155(4):1466–1475. 270. Davis BJ, Lennard DE, Lee CA, et al. Anovulation in cyclooxygenase-­2-­deficient mice is restored by prostaglandin E2 and interleukin-­lbeta. Endocrinology. 1999;140:2685–2695. 271. Reese J, Zhao X, Ma WG, et al. Comparative analysis of pharmacologic and/or genetic disruption of cyclooxygenase-­1 and cyclooxygenase-­2 function in female reproduction in mice. Endocrinology. 2001;142:3198–3206. 272. Mikuni M, Pall M, Peterson CM, et al. The selective prostaglandin endoperoxide synthase-­2 inhibitor, NS-­398, reduces prostaglandin production and ovulation in vivo and in vitro in the rat. Biol Reprod. 1998;59:1077–1083. 273. Pall M, Friden BE, Brannstrom M. Induction of delayed follicular rupture in the human by the selective COX-­2 inhibitor rofecoxib: a randomized double-­blind study. Hum Reprod. 2001;16:1323–1328. 274. Duffy DM, McGinnis LK, Vandevoort CA, et al. Mammalian oocytes are targets for prostaglandin E2 (PGE2) action. Reprod Biol Endocrinol. 2010;8:131. 275. Sugimoto Y, Yamasaki A, Segi E, et al. Failure of parturition in mice lacking the prostaglandin F receptor. Science. 1997;277:681–683. 276. Kennedy CR, Zhang Y, Brandon S, et al. Salt-­sensitive hypertension and reduced fertility in mice lacking the prostaglandin EP2 receptor. Nat Med. 1999;5:217–220. 277. Hizaki H, Segi E, Sugimoto Y, et al. Abortive expansion of the cumulus and impaired fertility in mice lacking the prostaglandin E receptor subtype EP(2). Proc Natl Acad Sci U S A. 1999;96:10501–10506. 278. Matsumoto H, Ma W, Smalley W, et al. Diversification of cyclooxygenase-­2-­derived prostaglandins in ovulation and implantation. Biol Reprod. 2001;64:1557–1565. 278A. Ben-­Ami I, Freimann S, Armon L, et al. PGE2 up-­regulates EGF-­ like growth factor biosynthesis in human granulosa cells: new insights into the coordination between PGE2 and LH in ovulation. Mol Hum Reprod. 2006;12(10):593–599. 278B. Waclawik A, Kaczynski P, Jabbour HN. Autocrine and paracrine mechanisms of prostaglandin E action on trophoblast/conceptus cells through the prostaglandin E receptor (PTGER2) during implantation. Endocrinology. 2013;154(10):3864–3876. 279. Markosyan N, Duffy DM. Prostaglandin E2 acts via multiple receptors to regulate plasminogen-­dependent proteolysis in the primate periovulatory follicle. Endocrinology. 2009;150(1):435–444. 280. Powell WS, Hammarström S, Samuelsson B, et al. Letter: prostaglandin-F2alpha receptor in human corpora lutea. Lancet. 1974;1:1120. 281. Bennegard B, Hahlin M, Wennberg E, et al. Local luteolytic effect of prostaglandin F2 alpha in the human corpus luteum. Fertil Steril. 1991;56:1070–1076. 282. Manaugh LC, Novy MJ. Effects of indomethacin on corpus luteum function and pregnancy in rhesus monkeys. Fertil Steril. 1976;27:588–598. 283. Sargent EL, Baughman WL, Novy MJ, et al. Intraluteal infusion of a prostaglandin synthesis inhibitor, sodium meclofenamate, causes premature luteolysis in rhesus monkeys. Endocrinology. 1988;123:2261–2269. 284. Huang JC, Wun WS, Goldsby JS, et al. Stimulation of embryo hatching and implantation by prostacyclin and peroxisome proliferator-­activated receptor delta activation: implication in IVF. Hum Reprod. 2007;22:807–814.

285. Kaczynski P, Kowalewski MP, Waclawik A. Prostaglandin F2α promotes angiogenesis and embryo-­maternal interactions during implantation. J Reprod Fertil. 2016;151(5):539–552. 286. Kaczynski P, Baryla M, Goryszewska E, Bauersachs S, Waclawik A. Prostaglandin F2α promotes embryo implantation and development in the pig. J Reprod Fertil. 2018;156(5):405–419. 287. Cheng JG, Stewart CL. Loss of cyclooxygenase-­2 retards decidual growth but does not inhibit embryo implantation or development to term. Biol Reprod. 2002;68:401–404. 288. Forman BM, Chen J, Evans RM. Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator-­activated receptors alpha and delta. Proc Natl Acad Sci U S A. 1997;94:4312–4317. 289. Murata T, Ushikubi F, Matsuoka T, et al. Altered pain perception and inflammatory response in mice lacking prostacyclin receptor. Nature. 1997;388:678–682. 290. Barak Y, Liao D, He W, et al. Effects of peroxisome proliferator-­ activated receptor delta on placentation, adiposity, and colorectal cancer. Proc Natl Acad Sci U S A. 2002;99:303–308. 291. Huang JC, Wun WS, Goldsby JS, et al. Prostacyclin receptor signaling and early embryo development in the mouse. Hum Reprod. 2007;22:2851–2856. 292. Koike H, Egawa H, Ohtsuka T, et al. Correlation between dysmenorrheic severity and prostaglandin production in women with endometriosis. Prostaglandins Leukot Essent Fatty Acids. 1992;46:133–137. 293. Dawood MY. Nonsteroidal anti-­inflammatory drugs and changing attitudes toward dysmenorrhea. Am J Med. 1988;84:23–29. 294. Hayes EC, Rock JA. COX-­2 inhibitors and their role in gynecology. Obstet Gynecol Surv. 2002;57:768–780. 295. Herington JL, O’Brien C, Robuck MF, et al. Prostaglandin-­endoperoxide synthase 1 mediates the timing of parturition in mice despite unhindered uterine contractility. Endocrinology. 2018;159(1):490–505. 296. Daniels SE, Talwalker S, Torri S, et al. Valdecoxib, a cyclooxygenase-­2-­specific inhibitor, is effective in treating primary dysmenorrhea. Obstet Gynecol. 2002;100:350–358. 297. Alfirevic Z, Keeney E, Dowswell T, et al. Labour induction with prostaglandins: a systematic review and network meta-­ analysis. BMJ. 2015;350:h217. 298. Lewis RB, Schulman JD. Influence of acetylsalicylic acid, an inhibitor of prostaglandin synthesis, on the duration of human gestation and labour. Lancet. 1973;2:1159–1161. 299. Skarnes RC, Harper MJ. Relationship between endotoxin-­induced abortion and the synthesis of prostaglandin F. Prostaglandins. 1972;1:191–203. 300. Fidel Jr PL, Romero R, Wolf N, et al. Systemic and local cytokine profiles in endotoxin-­induced preterm parturition in mice. Am J Obstet Gynecol. 1994;170:1467–1475. 301. Olson DM, Ammann C. Role of the prostaglandins in labour and prostaglandin receptor inhibitors in the prevention of preterm labour. Front Biosci. 2007;12:1329–1343. 302. Gross GA, Imamura T, Luedke C, et al. Opposing actions of prostaglandins and oxytocin determine the onset of murine labor. Proc Natl Acad Sci U S A. 1998;95:11875–11879. 303. Yu Y, Cheng Y, Fan J, et al. Differential impact of prostaglandin H synthase 1 knockdown on platelets and parturition. J Clin Invest. 2005;115:986–995. 304. Cook JL, Shallow MC, Zaragoza DB, et al. Mouse placental prostaglandins are associated with uterine activation and the timing of birth. Biol Reprod. 2002;68:579–587. 305. Winchester SK, Imamura T, Gross GA, et al. Coordinate regulation of prostaglandin metabolism for induction of parturition in mice. Endocrinology. 2002;143:2593–2598. 306. Li X, Ballantyne LL, Crawford MC, FitzGerald GA, Funk CD. Isoform-­specific compensation of cyclooxygenase (ptgs) genes during implantation and late-­stage pregnancy. Sci Rep. 2018;8(1):12097. 307. Durn JH, Marshall KM, Farrar D, et al. Lipidomic analysis reveals prostanoid profiles in human term pregnant myometrium. Prostaglandins Leukot Essent Fatty Acids. 2010;82(1):21–26. 308. Fetalvero KM, Zhang P, Shyu M, et al. Prostacyclin primes pregnant human myometrium for an enhanced contractile response in parturition. J Clin Invest. 2008;118(12):3966–3979. 309. Taggart MJ, Europe-­Finner GN, Mitchell BF. Possible dual roles for prostacyclin in human pregnancy and labor. J Clin Invest. 2008;118(12):3829–3832. al. The roles of prostaglandin EP 1 and 3 receptors in the control of human myometrial contractility. J Clin Endocrinol Metab. 2012;97(2):489–498.

CHAPTER 4  Steroid Hormones and Other Lipid Molecules Involved in Human Reproduction 311. Astle S, Newton R, Thornton S, et al. Expression and regulation of prostaglandin E synthase isoforms in human myometrium with labour. Mol Hum Reprod. 2007;13(1):69–75. 312. Mozurkewich EL, Chilimigras JL, Berman DR, et al. Methods of induction of labour: a systematic review. BMC Pregnancy Childbirth. 2011;11:84. 313. Gross G, Imamura T, Vogt SK, et al. Inhibition of cyclooxygenase-­2 prevents inflammation-­mediated preterm labor in the mouse. Am J Physiol Regul Integr Comp Physiol. 2000;278:1415–1423. 314. Vermillion ST, Landen CN. Prostaglandin inhibitors as tocolytic agents. Semin Perinatol. 2001;25:256–262. 315. Stika CS, Gross GA, Leguizamon G, et al. A prospective randomized safety trial of celecoxib for treatment of preterm labor. Am J Obstet Gynecol. 2002;187:653–660. 316. Loftin CD, Trivedi DB, Langenbach R. Cyclooxygenase-­1-­ selective inhibition prolongs gestation in mice without adverse effects on the ductus arteriosus. J Clin Invest. 2002;110:549–557. 317. Takahashi Y, Roman C, Chemtob S, et al. Cyclooxygenase-­ 2 inhibitors constrict the fetal lamb ductus arteriosus both in vitro and in vivo. Am J Physiol Regul Integr Comp Physiol. 2000;278:R1496–R1505. 318. Smith GC. The pharmacology of the ductus arteriosus. Pharmacol Rev. 1998;50:35–58. 319. Momma K, Toyoshima K, Takeuchi D, Imamura S, Nakanishi T. In vivo constriction of the fetal and neonatal ductus arteriosus by a prostanoid EP4-­receptor antagonist in rats. Pediatr Res. 2005;58(5):971–975. 320. Sakuma T, Akaike T, Minamisawa S. Prostaglandin E2 receptor EP4 inhibition contracts rat ductus arteriosus. Circ J. 2018;83(1):209–216. 321. Nguyen M, Camenisch T, Snouwaert JN, et al. The prostaglandin receptor EP4 triggers remodeling of the cardiovascular system at birth. Nature. 1997;390:78–81. 322. Coggins KG, Latour A, Nguyen MS, et al. Metabolism of PGE2 by prostaglandin dehydrogenase is essential for remodeling the ductus arteriosus. Nat Med. 2002;8:91–92. 323. Yokoyama U, Minamisawa S, Quan H, et al. Chronic activation of the prostaglandin receptor EP4 promotes hyaluronan-­mediated neointimal formation in the ductus arteriosus. J Clin Invest. 2006;16:3026–3034. 324. Reese J, Paria BC, Brown N, et al. Coordinated regulation of fetal and maternal prostaglandins directs successful birth and postnatal adaptation in the mouse. Proc Natl Acad Sci U S A. 2000;97:9759–9764. 325. Loftin CD, Trivedi DB, Tiano HF, et al. Failure of ductus arteriosus closure and remodeling in neonatal mice deficient in cyclooxygenase-­1 and cyclooxygenase-­2. Proc Natl Acad Sci U S A. 2001;98:1059–1064. 326. Segi E, Sugimoto Y, Yamasaki A, et al. Patent ductus arteriosus and neonatal death in prostaglandin receptor EP4-­deficient mice. Biochem Biophys Res Commun. 1998;246:7–12. 327. Fitzgerald DJ, Mayo G, Catella F, et al. Increased thromboxane biosynthesis in normal pregnancy is mainly derived from platelets. Am J Obstet Gynecol. 1987;157:325–330. 328. Fitzgerald DJ, Rocki W, Murray R, et al. Thromboxane A2 synthesis in pregnancy-­induced hypertension. Lancet. 1990;335:751–754. 329. Rocca B, Loeb AL, Strauss III JF, et al. Directed vascular expression of the thromboxane A2 receptor results in intrauterine growth retardation. Nat Med. 2000;6:219–221. 330. Rolnik DL, Wright D, Poon LC, et al. Aspirin versus placebo in pregnancies at high risk for preterm preeclampsia. N Engl J Med. 2017;377(7):613–622. 331. Wallenburg HC. Prevention of pre-­eclampsia: status and perspectives 2000. Eur J Obstet Gynecol Reprod Biol. 2001;94:13–22. 332. Caritis S, Sibai B, Hauth J, et al. Low-­dose aspirin to prevent preeclampsia in women at high risk. N Engl J Med. 1998;338:701–705. 333. Mills JL, DerSimonian R, Raymond E, et al. Prostacyclin and thromboxane changes predating clinical onset of preeclampsia: a multicenter prospective study. JAMA. 1999;282:356–362. 334. Henderson JT, Whitlock EP, O’Connor E, et al. Low-­dose aspirin for prevention of morbidity and mortality from preeclampsia: a systematic evidence review for the U.S. Preventive Services Task Force. Ann Intern Med. 2014;160(10):695–703. 335. Tolcher MC, Chu DM, Hollier LM, et al. Impact of USPSTF recommendations for aspirin for prevention of recurrent preeclamp sia. Am J Obstet Gynecol

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336. McKanna JA, Zhang MZ, Wang JL, et al. Constitutive expression of cyclooxygenase-­ 2 in rat vas deferens. Am J Physiol. 1998;275:R227–R233. 337. Cheuk BL, Leung PS, Lo AC, et al. Androgen control of cyclooxygenase expression in the rat epididymis. Biol Reprod. 2000;63:775–780. 338. Kirschenbaum A, Liotta DR, Yao S, et al. Immunohistochemical localization of cyclooxygenase-­ 1 and cyclooxygenase-­ 2 in the human fetal and adult male reproductive tracts. J Clin Endocrinol Metab. 2000;85:3436–3441. 339. Lazarus M, Munday CJ, Eguchi N, et al. Immunohistochemical localization of microsomal PGE synthase-­1 and cyclooxygenases in male mouse reproductive organs. Endocrinology. 2002;143:2410–2419. 340. Walch L, Morris PL. Cyclooxygenase 2 pathway mediates IL-­ 1beta regulation of IL-­1alpha, -­Ibeta, and IL-­6 mRNA levels in Leydig cell progenitors. Endocrinology. 2002;143:3276–3283. 341. Cheuk BL, Chew SB, Fiscus RR, et al. Cyclooxygenase-­2 regulates apoptosis in rat epididymis through prostaglandin D2. Biol Reprod. 2002;66:374–380. 342. Alexandre B, Lemaire A, Desvaux P, et al. Intracavernous injections of prostaglandin E1 for erectile dysfunction: patient satisfaction and quality of sex life on long-­term treatment. J Sex Med. 2007;4(2):426–431. 343. Khan MA, Thompson CS, Sullivan ME, et al. The role of prostaglandins in the aetiology and treatment of erectile dysfunction. Prostaglandins Leukot Essent Fatty Acids. 1999;60:169–174. 344. Edmonds JW, Prasain JK, Dorand D, et al. Insulin/FOXO signaling regulates ovarian prostaglandins critical for reproduction. Dev Cell. 2010;19(6):858–871. 345. Yousef MS, Marey MA, Hambruch N, Hayakawa H, Shimizu T, Hussien HA, Abdel-Razek AK, Pfarrer C, Miyamoto A. Sperm Binding to Oviduct Epithelial Cells Enhances TGFB1 and IL10 Expressions in Epithelial Cells as Well as Neutrophils In Vitro: Prostaglandin E2 As a Main Regulator of Anti-Inflammatory Response in the Bovine Oviduct. PLoS One. 2016;11(9): e0162309. 346. Ye X. Lysophospholipid signaling in the function and pathology of the reproductive system. Hum Reprod Update. 2008;14(5):519–536. 347. Guo L, Ou X, Li H, et al. Roles of sphingosine-­1-­phosphate in reproduction. Reprod Sci. 2014;21(5):550–554. 348. Budnik LT, Mukhopadhyay AK. Lysophosphatidic acid and its role in reproduction. Biol Reprod. 2002;66:859–865. 349. Aoki J. Mechanisms of lysophosphatidic acid production. Semin Cell Dev Biol. 2004;15:477–489. 350. Chun J, Hla T, Lynch KR, et al. International union of basic and clinical pharmacology. LXXVIII. Lysophospholipid receptor nomenclature. Pharmacol Rev. 2010;62(4):579–587. 351. Hla T, Lee MJ, Ancellin N, et al. Lysophospholipids: receptor revelations. Science. 2001;294:1875–1878. 352. Spiegel S, Milstien S. The outs and the ins of sphingosine-­1-­ phosphate in immunity. Nat Rev Immunol. 2011;11(6):403–415. 353. Hla T, Brinkmann V. Sphingosine 1-­phosphate (S1P): physiology and the effects of S1P receptor modulation. Neurology. 2011;76(8 suppl 3):S3–S8. 354. Sanchez T, Hla T. Structural and functional characteristics of S1P receptors. J Cell Biochem. 2004;92:913–922. 355. Tokumura A, Fukuzawa K, Yamada S, et al. Stimulatory effect of lysophosphatidic acids on uterine smooth muscles of non-­pregnant rats. Arch Int Pharmacodyn Ther. 1980;245:74–83. 356. Ye X, Hama K, Contos JJ, et al. LPA3-­mediated lysophosphatidic acid signalling in embryo implantation and spacing. Nature. 2005;435:104–108. 357. Hama K, Aoki J, Bandoh K, et al. Lysophosphatidic receptor, LPA3, is positively and negatively regulated by progesterone and estrogen in the mouse uterus. Life Sci. 2006;79:1736–1740. 358. Kennedy TG. Evidence for a role for prostaglandins in the initiation of blastocyst implantation in the rat. Biol Reprod. 1977;16:286–291. 359. Kinoshita K, Satoh K, Ishihara O, et al. Involvement of prostaglandins in implantation in the pregnant mouse. Adv Prostaglandin Thromboxane Leukot Res. 1985;15:605–607. 360. Song H, Lim H, Paria BC, et al. Cytosolic phospholipase A2alpha is crucial [correction of A2alpha deficiency is crucial] for ‘on-­ time’ embryo implantation that directs subsequent development. Development. 2002;129(12):2879–2889. 361. Ye X, Skinner MK, Kennedy G, et al. Age-­dependent loss of sperm production in mice via impaired lysophosphatidic acid signaling. Biol Reprod

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109.e8 PART I  The Fundamentals of Reproduction 362. Yanagida K, Ishii S. Non-­Edg family LPA receptors: the cutting edge of LPA research. J Biochem. 2011;150(3):223–232. 363. Skaznik-­Wikiel ME, Kaneko-­ Tarui T, Kashiwagi A, et al. Sphingosine-­ 1-­ phosphate receptor expression and signaling correlate with uterine prostaglandin-­ endoperoxide synthase 2 expression and angiogenesis during early pregnancy. Biol Reprod. 2006;74:569–576. 364. Tilly JL. Commuting the death sentence: how oocytes strive to survive. Nat Rev Mol Cell Biol. 2001;2:838–848.

365. Perez GI, Jurisicova A, Matikainen T, et al. A central role for ceramide in the age-­related acceleration of apoptosis in the female germline. FASEB J. 2005;19:860–862. 366. Suomalainen L, Hakala JK, Pentikainen V, et al. Sphingosine-­1-­ phosphate in inhibition of male germ cell apoptosis in the human testis. J Clin Endocrinol Metab. 2003;88:5572–5579. 367. Otala M, Pentikainen MO, Matikainen T, et al. Effects of acid sphingomyelinase deficiency on male germ cell development and programmed cell death. Biol Reprod. 2005;72:86–96.

5

Steroid Hormone Action Shannon Whirledge and John A. Cidlowski

OUTLINE STEROID HORMONE RECEPTORS ACT AS LIGAND-­ DEPENDENT TRANSCRIPTION ACTIVATORS OR REPRESSORS STEROID HORMONE RECEPTOR STRUCTURE AND THE EVOLUTION OF SPECIFICITY STEROID HORMONE RECEPTOR FUNCTION Estrogen Receptor Progesterone Receptor Androgen Receptor Glucocorticoid Receptor Mineralocorticoid Receptor GENERAL FACTORS THAT INFLUENCE STEROID HORMONE ACTION Hormone Bioavailability Receptor Expression Ligand-­Bound Changes to Receptor Conformation Posttranslational Modifications of the Steroid Hormone Receptors Interaction With DNA Interaction With Coactivators and Corepressors Interaction With Other Transcription Factors Nongenomic Actions of Steroids Signaling via Second Messenger Cascades SUMMARY

STEROID HORMONE RECEPTORS ACT AS LIGAND-­ DEPENDENT TRANSCRIPTION ACTIVATORS OR REPRESSORS

• S  teroid hormones are derived from the metabolic conversion of cholesterol into biologically active steroid products that bind to intracellular receptors with high specificity. • The canonical mechanism of action for the steroid hormone receptors involves regulating gene transcription through transactivation and transrepression. • Precise regulation of gene transcription is essential for development, physiology, and homeostasis.    Steroids are small, lipophilic hormones synthesized from a common precursor molecule, cholesterol, through a complex biosynthetic process in specific tissues and glands throughout the body (see Chapter 4). Despite their shared molecular origin and basic structural similarities, the steroids mineralocorticoids, glucocorticoids, estrogens, progestins, and androgens are distinct classes of hormones that interact with specific, high-­ affinity receptors to exert their unique biological effects (mineralocorticoid [MR], glucocorticoid [GR], estrogen [ER], progestin [PR], and androgen [AR]). These hormones control diverse physiological and cellular processes and affect almost all aspects of eukaryotic

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physiology, from sexual differentiation, growth, and reproductive functions to immunity, metabolism, behavior, and learning.1 Consequently, a clear and complete understanding of the general mechanisms of steroid hormone action, as well as those activities that occur in a tissue-­and cell-­type-­specific manner, is of critical importance for the promotion of health and the understanding of disease processes. This chapter reviews the structural similarities and differences among the steroid hormone receptors, as well as what is known regarding their mechanisms of action. The classic mode of action entails simple diffusion of steroid hormones into the cell, where they interact with cognate receptors and stimulate or inhibit transcription of target genes (Fig. 5.1).2 The hormone-­dependent changes in receptor conformation can drive both the transactivation and transrepression of gene expression by (1) altering interactions with molecular chaperones that keep the receptor in a ligand-­independent state; (2) inducing posttranslational modifications that alter receptor activity; (3) promoting the formation of receptor dimers; (4) enhancing interactions with specific DNA sequences (hormone response elements); and (5) facilitating recruitment of coactivator or corepressor proteins that alter chromatin structure and contact with the basal transcription machinery.3 Recent mechanistic studies have built on the classic mode of action to reveal a complex regulatory network of interacting factors and chromatin state.4,5 For example, steroid hormone receptors have been shown to interact with closed chromatin through the recognition of partial DNA sequence motifs to enable other transcription factors to engage chromatin and form regulatory complexes (e.g., pioneer factors) or work through receptor cooperation to initiate the opening of chromatin and allow for the binding of other steroid hormone receptors or secondary transcription factors.6,7 In addition to reviewing the various mechanisms of steroid hormone action, this chapter will discuss factors that regulate hormone activity, as well as nonclassical modes of action for steroid hormones and their receptors.

STEROID HORMONE RECEPTOR STRUCTURE AND THE EVOLUTION OF SPECIFICITY • T  he steroid hormone receptors belong to a large family of transcription factors called nuclear receptors. • The structure of the steroid hormone receptors is modular, with distinct domains. The steroid hormone receptors contain a highly conserved DNA-­binding domain (DBD), a moderately conserved ligand-­binding domain (LBD), and less well-­conserved amino-­ and carboxy-­terminal domains. • Phylogenetic analysis determined that the steroid hormone receptors cluster separately from other ligand-­dependent transcription factors. The evolution of the steroid hormone receptors is controversial and may reflect gene duplication, mutation, and functional divergence.    The steroid hormone receptors belong to a larger family of structurally and evolutionarily related proteins called nuclear receptors, encoded by 48 genes in the human genome.8–10 All nuclear receptors, including the steroid hormone receptors, exhibit a modular structure composed of distinct domains (Fig. 5.2).

CHAPTER 5  Steroid Hormone Action

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Steroids

5

OH

Molecular chaperone complex

HSP

HO

SR

OH

conformational change

LBP

HSP

Cytoplasm

Dissociation of chaperones P HSP

HSP

SR

Hormone binding and

HSP

HO

Receptor phosphorylation

SR

Plasma membrane

HSP

OH

HO

OH

HO

SR

Basal transcription machinery

Histone acetylase

SR TF

HO

Nucleus

A

Transcription initiation

RE

HRE

A P

P SR

Histone acetylase

OH

HO

Histones

Coactivators

OH

SR

A

Acetylation

P

HSP

Nuclear translocation

Receptor dimerization

HSP

A

Acetylation

Coactivators

Basal transcription machinery

Histones

Transcription initiation

OH HO

Sp1 Sp1 site P

P SR

SR

OH

HO

Histone deacetylase

Deacetylation

Corepressors Histones

OH HO

Transcriptional repression

HRE P SR Transcriptional repression

TF

HO

OH

RE

HRE P

P SR

SR

OH

HO

OH HO

p65 p50

NF-κB site

Basal transcription machinery Transcriptional repression

Fig. 5.1 General mechanism of action for cytoplasmic steroid receptors as described in the text.  The two subunits of nuclear factor kappa B are p50 and p65. A, Acetyl group; HRE, hormone response element; HSP, heat shock protein; LBP, ligand-­binding pocket; NF, nuclear factor; P, phosphate group; SR, steroid receptor.

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PART I  The Fundamentals of Reproduction Steroid receptor domains NH-

A/B

C

AF1

D

E

DNA Hinge binding

F -COOH

Ligand AF2 binding

Human steroid receptor OH

ERα 1

180

263 302

552 595

H H

H HO

ERβ

Estradiol 1

144

227 255

391

470

504 530

PRA 1

524

770

O

H

PRB 1

AF3

556

632

687

H

H

933 O

Progesterone PRC 1

37

92

338 OH H

AR 1

558

624

676

919

H

H O

Testosterone O

GRα

HO

1

421

486

528

OH OH

777

H H

H O

Cortisol

GRβ 1

421

486

528

O

742

O

OH

HO H H

MR

H

O

1

602

670

734

984

Aldosterone

Fig. 5.2 Schematic diagram of the primary structure of a generic steroid receptor and its functional domains.  Region A/B contains transactivation function 1 (AF1) domain. Region C contains the DNA-­ binding domain (DBD). Region D is the hinge region. Region E contains the ligand-­binding domain (LBD). Region F contains the transactivation function 2 (AF2) domain. The primary structure of human steroid receptors, their isoforms, and their physiological ligands: AR, androgen receptor; ER, estrogen receptor; GR, glucocorticoid receptor; MR, mineralocorticoid receptor; PRA, progesterone receptor isoform A; PRB, progesterone receptor isoform B; PRC, progesterone receptor isoform C.

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Genome-wide gene duplication in jawed vertebrates

5

PR 3-Ketogonadal steroid receptor AR

3-Ketosteroid receptor

Lamprey PR

Corticoid receptor

GR

Ancestral ER

MR Lamprey CR ERα

ERβ Lamprey ER Estrogen-related receptor

In general, nuclear receptors contain a variable amino-­terminal region (A/B), a highly conserved DBD (C), a highly variable hinge region (D), and a moderately conserved hormone-­or ligand-­binding domain (LBD) (E).11,12 Some receptors also contain a carboxy-­terminal F domain. The primary structure of each human steroid hormone receptor is shown, along with its physiological ligand (Fig. 5.2). Specific residues within the DBD and LBD play an important part in receptor dimerization, which is critical because most nuclear receptors are only transcriptionally active as homo-­or heterodimers. Finally, nuclear receptors have regions called activation function 1 and 2 (AF1 and AF2) that are required to transactivate gene expression.13 Whereas the activity of AF1 is usually ligand-­independent and located in the A/B domain, AF2 is found in the LBD and is predominantly regulated by hormone binding. Some nuclear receptors have defined natural ligands, such as the steroid hormones, thyroid hormones, retinoids, or vitamin D, but the ligand for other nuclear receptors has not yet been identified and are therefore termed “orphan receptors.” The finding that diverse compounds act as ligands for nuclear receptors and that some receptors have no apparent ligand led to the hypothesis that ancestral nuclear receptors were constitutive transcription factors that independently evolved the ability to bind ligands.14,15 However, a second hypothesis posits that ancestral nuclear receptors were ligand-­dependent transcription factors that evolved specificity for different ligands by gene duplication, mutation, and functional divergence. Under this hypothesis, the yet-­to-­be-­identified ligands for the orphan receptors are likely intermediates in the synthesis of ligands for related receptors.16 There are several lines of evidence that favors the latter hypothesis for the evolution of ligand binding in the steroid hormone receptor family. For example, the primary,

Fig. 5.3 Phylogeny of the steroid receptor gene family.  AR, Androgen receptor; CR, corticoid receptor; ER, estrogen receptor; GR, glucocorticoid receptor; MR, mineralocorticoid receptor; PR, progestin receptor.

secondary, and tertiary structures of the LBD from different steroid hormone receptors are highly similar.17–20 Furthermore, detailed sequence, structural, and functional analyses support the hypothesis that the ancestral steroid hormone receptor-­bound estrogens and specificity for other steroids evolved by serial and parallel duplications of the ancestral gene, mutation of nucleotides coding for specific amino acids, and structural and functional divergence of the paralogs.16,21,22 Indeed, reconstructing ancestral sequences, using a maximum likelihood approach, suggests that the first steroid hormone receptor was an ER-­like molecule (Fig. 5.3).16 When this gene was duplicated, one copy was constrained by natural selection and retained its function as an estrogen-­binding receptor, while the other copy evolved specificity for 3-­ketosteroid-­like ligands (Fig. 5.3). Duplication of the latter gene then produced a corticoid receptor-­like protein and a receptor for 3-­ketogonadal steroid-­like molecules (e.g., androgens, progestins, or both). This final duplication then led to the evolution of the true androgen and progesterone receptors from the ancestral 3-­ketogonadal steroid hormone receptor. Crystal structures of the various steroid hormone receptors have shown that the LBD folds into a highly homologous three-­layered structure with a small ligand-­binding pocket in the center, while domain swapping of the entire LBD between steroid hormone receptors has shown that this region determines specificity for particular classes of steroid hormones. The ligand-­binding pocket is composed of roughly 30 amino acids that are in close proximity or make direct contact with hormones when bound to their cognate receptors. In agreement with structural studies, experiments using site-­directed mutagenesis indicate that specific, but minor, changes of binding pocket can lead to

114

PART I  The Fundamentals of Reproduction C G H I S Y K R A G R D E V K I D L S T N I C C C C ND P G V E R 2 + 2 + Zn Zn A S L G GA C C C C KV FFKRAVEGQHNYL PKL RYRKCLQA A

Fig. 5.4 Amino acid sequence of the DNA-­binding domain of human glucocorticoid receptor showing two zinc fingers and the P box, which is outlined.  The three mutant residues that change glucocorticoid receptor binding specificity from glucocorticoid-­ responsive element and estrogen-­responsive element are indicated by the arrows.

dramatic changes in the binding specificity of the steroid hormone receptors. For example, the human PR, GR, and MR contain a conserved cysteine residue that appears to be critical for contacting the C20 keto group found in progestins, glucocorticoids, and mineralocorticoids.16,23 Mutation of the corresponding threonine to cysteine in the AR reduces its affinity for androgens and allows the receptor to transactivate in the presence of progesterone and corticoids.24 Based on structure-­function studies of this sort and phylogenetic analyses, Thornton16 proposed a series of relatively minor amino acid changes that may account for broad changes in hormone specificity during the evolution of the steroid hormone receptors. Similar studies of the DBD have defined the molecular basis for interactions between steroid hormone receptors and particular DNA sequences.25,26 The DBD of nuclear receptors contains two zinc fingers. The first zinc finger interacts with the major groove of DNA, whereas the second is involved in receptor dimerization. Mutation of three residues within a five-­residue motif known as the Proximal box (P box) of the first zinc finger of the GR to the corresponding residues in the ER changes the binding specificity from DNA sequences called glucocorticoid-­response elements (GREs) to estrogen-­ response elements (EREs) and vice versa (Fig. 5.4). Although the DBD is very highly conserved among nuclear receptors, the hormone response elements are variable among different receptors, which may in part account for receptor-­specific regulation of distinct sets of genes. These examples illustrate how site-­directed mutagenesis and fine-­scale comparison of amino acid sequences among receptors, in the context of the tertiary structure of the DBD and LBD, facilitate testable hypotheses regarding the evolution of signaling and regulation of gene expression by different classes of steroids.27

STEROID HORMONE RECEPTOR FUNCTION • T  he steroid hormone receptors have well-­described functions in the reproductive tract but also critically regulate general physiology. • Transgenic animal models and mutations identified in humans have provided insight into the cell-­specific activities of the steroid hormones and their receptors. • ER, PR, and AR are indispensable for reproductive function but also regulate aspects of physiology outside of the reproductive system.

• G  R signaling is essential for life after birth, maintaining general physiologic homeostasis, and integrating the hypothalamic-­pituitary-­ adrenal (HPA) axis with reproductive functions mediated by the hypothalamic-­pituitary-­gonadal (HPG) axis. • MR is required for electrolyte balance and fluid transport, where insufficiency or deficiency can lead to mortality.    Steroid hormone receptors have taken on distinct physiological roles during evolution, which has been demonstrated by transgenic animal models and case reports in humans with genetic mutations/deletions. For example, the sex hormone receptors demonstrate a pattern of expression primarily restricted to cells and organs of the HPG axis and play an essential role in sexual differentiation and reproduction. However, these steroid hormone receptors are not sex-­limited and also have some functions outside of the reproductive system in both sexes. In contrast, the GR is expressed ubiquitously and plays a role in a wide range of physiological processes, including, development, immune function, cognition and behavior, cardiovascular health, metabolic homeostasis, and reproduction.28,29 Outside of their roles in reproductive functions and overall physiology, the steroid hormone receptors can drive disease pathology when signaling becomes unregulated. Functions of ER, PR, AR, GR, and MR in physiology and pathophysiology are described briefly below.

Estrogen Receptor The biological effects of estrogens were originally believed to be mediated by a single receptor (ERα) until the cloning of a second receptor (ERβ) and reports of estrogen binding to a membrane-­ bound G protein-­ coupled receptor (GPR30/GPER) several decades later.30,31 The estrogen receptors are products of different genes and demonstrate distinct expression patterns across tissues and cell types (Table 5.1).32 Along with their tissue type-­ specific distribution, the different estrogen receptors regulate unique biological functions, which have been characterized in transgenic animal models. For example, the ovarian phenotypes of ERα and ERβ knockout mice are distinct, likely due to the cellular distribution of these two receptors in the ovary, where ovarian granulosa cells express ERβ, and ERα is primarily found in the germinal epithelial cells, interstitial cells, and theca cells of the ovary.33–­35 Subsequently, the ERα knockout mouse is anovulatory and accumulates cystic and hemorrhagic follicles, whereas the ERβ knockout mouse contains histologically normal ovaries but still displays impaired ovulation.36 In the uterus, the absence of ERα results in a significant infertile phenotype due to a hypoplastic uterus and defects in implantation and decidualization; however, the loss of ERβ does not alter uterine development, function, or cellular responses to estradiol.37,38 Subsequently, cell-­specific deletion models have dissected the biological effects of ERα in the uterine stroma and epithelial cells.39–41 The mammary gland is also a key organ for ERα signaling. ERα knockout mice demonstrate impaired ductal growth and differentiation following the onset of endogenous estrogen production during puberty.42,43 The loss of ERβ does not alter ductal growth but appears to play a role in differentiation of the mammary gland during pregnancy.44 In males, loss of ERα results in infertility related to defects in fluid resorption in the efferent ducts and epididymis with progressive deterioration of testicular morphology, while the ERβ knockout males are fertile with phenotypically normal testis.45,46 Gene targeting technology has also led to the creation of mouse models with mutations in the functional domains of ERα, which have allowed the functional domains to be associated with specific physiological functions.47–49 For example, the functional domain mediating the metabolic functions of ERα have been discriminated using mouse models that selectively target the AF1 and AF2 domains.49,50

CHAPTER 5  Steroid Hormone Action

TABLE 5.1  Tissue-­Specific Patterns of ERα and ERβ mRNA Expression in the Rat Receptor* Tissue

ERα

ERβ

Epididymis Prostate Testis Pituitary Ovary Uterus Bladder Lung Liver Kidney Thymus Adrenal Olfactory lobe Cerebellum Brain stem Spinal cord Heart

+++ + +++ ++ +++ +++ + 0 + ++ + ++ 0 0 0 0 +

+ +++ + + +++ ++ ++ + 0 0 + 0 + + + + 0

*Relative levels of expression are indicated by the number of plus signs: 0, not detected; +, low; ++, medium; +++, high. (From Kuiper GGJM, Carlsson B, Grandien K, et al. Comparison of the ligand binding specificity and transcript tissue distribution of estrogen receptors α and β. Endocrinology. 1997;138:863–870.)

TABLE 5.2  Binding Affinity of Various Ligands for ERα and ERβ Relative to Binding Affinity of E2 Relative Binding Affinity* Ligand

ERα

ERβ

E2 Diethylstilbestrol Hexestrol Dienestrol Estrone 17α-­Estradiol Moxestrol Estriol 4-­OH-­Estradiol 2-­OH-­Estradiol Estrone-­3-­sulfate 4-­OH-­Tamoxifen ICI-­164384 Nafoxidine Clomifene Tamoxifen Coumestrol Genestein Bisphenol A Methoxychlor

100 468 302 223 60 58 43 14 13 7 G (Thr685Arg) were identified.27 These variants occur in DNA-­binding regions of the protein and are likely to reduce PRDM9 function, but this alteration has not been evaluated. In mice, PRDM9 requirement and modifier function is also sexually dimorphic. Male mice engineered to express a point mutation in the PRDM9 PR/SET domain required for methyltransferase activity are sterile, whereas female mice with this mutation are fertile.28 Therefore, both female mice and women can use a PRDM9-­independent pathway to repair DNA. DSBs then elicit downstream DNA repair mechanisms, which begin with homologous chromosome pairing, a process of chromosome movement that allows homologs to find one another. These movements end in synapsis, a mechanism that tethers the homologs together. These processes initiate in leptonema and continue to develop in zygonema. Completion of synapsis, through the formation of tripartite structures called the synaptonemal complex (SC) on every homologous pair of chromosomes, marks the pachynema stage. When SC formation begins, axial elements accumulate along the length of the homologs and provide docking sites for proteins that control recombination and cohesion. After deposition of two transverse filaments, the homologs are fully synapsed (Fig. 9.4). The building of the SC is critical to successful meiosis. Mutations in genes that encode SC proteins confer female infertility, typically causing recurrent miscarriages or primary ovarian insufficiency.29,30 For example, sequencing of SC component SYCP3 in women who presented with recurrent miscarriages identified a gene variant (NM_153694.1:c.657T>C) that alters SYCP3 splicing and its ability to modify DNA structures in vitro.31,32 In addition, microdeletions and point mutations in SC component SYCE1 are associated with primary ovarian insufficiency.33,34 The point mutation (NM_130784.2:c.613C>T) has subsequently been assessed in a knock-­in mouse model and these animals also lack follicles and oocytes.35 While homologs are synapsing, homologous chromosome recombination is also occurring—these processes are dependent upon one another. While chromosomes are unsynapsed,

9

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DSBs form Chromosomes condense Chromosomes DNA synthesis begin to pair Interphase Leptonema

Zygonema

RAD51/DMC1

Synapsis is completed Crossover occurs

Chromosomes desynapse

Pachynema

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MSH4/MSH5

HORMAD1

Cohesin core

HORMAD2

Chromatin loop

Chromosomes condense Chiasma visible Metaphase I

MLH1/MLH3

SYCP2/3

or if they fail to synapse, HORMA domain containing 1 and 2 (HORMAD1/2) proteins are deposited to unsynapsed areas. But, once synapsed, HORMADs must be removed. This removal is achieved through thyroid hormone receptor interactor 13 (TRIP13 or PCH2 in lower eukaryotes), an AAA-­ATPase that participates in noncrossover DSB repair (see below). Variants in TRIP13 are linked with female infertility phenotypes both in mice and humans. Female Trip13-­knockout mice are sterile and have premature ovarian insufficiency (POI).36 However, the phenotype of women harboring missense Trip13 variants is different because these women have oocytes. These oocytes instead fail to undergo meiotic maturation,37 indicating additional unknown functions of this protein during later stages of meiosis. In males, the presence of nonhomologous X and Y chromosomes poses an interesting pairing conundrum. The mechanism by which spermatocytes tackle this problem is by creating an XY (or sex) body, where markers of asynapsis persist because of limited homology to enable pairing. Homologous recombination only occurs in the small homologous region, called the pseudoautosomal region,38 and transcription is silenced through chromatin modifications. If silencing, called meiotic sex chromosome inactivation (MSCI), fails to occur, germ cells arrest in pachytene likely because of aberrant gene expression incompatible with meiotic progression.39 Interestingly, when XO female mice are evaluated for MSCI, the results indicate mosaicism; some oocytes arrest whereas others will complete prophase I.40 It is unclear why MSCI is more robust in male mice at this time, but in sex-­reversed XY females (oocytes have an X and Y), mosaic silencing also occurs. Therefore, MSCI is not conferred by the Y chromosome but is dependent upon the sex of the gonad. DSBs are genotoxic and DNA repair is therefore essential for successful meiosis. During the repair process, resected DNA becomes single-­stranded, and DNA helicases like MCM8/9 protect the vulnerable DNA. Both male and female Mcm8-­knockout mice are sterile and Mcm9 knockout female mice are sterile.41 Human genome sequencing studies have since identified variants in Mcm8/9 associated with POI42; when one of the truncated MCM9 mutants was evaluated in a heterologous DNA repair system, it failed to localize to sites of DNA damage. In

SYCP1 SYCE1−3/TEX12

Fig. 9.4 Schematic representation of the events occurring between homologous chromosomes during prophase of meiosis I.  Substages of prophase I and relative progression of synapsis and recombination are depicted with spatiotemporal distribution of proteins involved in the synaptonemal complex formation and recombination. (Modified from Bolcun-­Filas E, Schimenti JC. Genetics of meiosis and recombination in mice. Int Rev Cell Mol Biol. 2012;298:179–227.)

mice, if chromosomes fail to synapse or repair, checkpoints exist to remove these oocytes from the ovary, also called the ovarian reserve. The DNA damage checkpoint, involving signaling via the checkpoint kinase 1 (CHEK1) and CHEK2, detects persistent DSBs and unsynapsed chromosomes and triggers transformation-­ related protein 53 (TRP53)/ transformation related protein 63 (TRP63)-­dependent apoptosis.43–46 In mouse spermatocytes, a similar pathway exists to eliminate germ cells with autosomal asynapsis.47 In human males with severe azoospermia, a meiotic arrest is also observed. Testes from these men show either XY body formation failure (i.e., no MSCI) or activated DNA damage checkpoint pathways.48 There are two outcomes of recombination-­based DNA repair (Fig. 9.5): 1) crossovers where the homologous chromosome serves as a repair template and results in a recombinant chromosome, one distinct from either parent chromosome or 2) noncrossovers where there is genetic exchange from the sister or the homolog causing gene conversion, but the sequence surrounding the repaired site remains unchanged. If DNA is repaired via crossing over, a physical linkage of the homologs occurs. This linkage, called a chiasma, keeps homologous chromosomes together until chromosome segregation in MI. Therefore at least one chiasma is required per chromosome for accurate separation of homologs in MI. Several genes are required for crossover recombination and fertility in mice.49–53 Mismatch repair proteins MLH1 and MLH3 (mutL homologs 1 and 3) resolve recombination intermediates and MLH3 variants are associated with human infertility in both males54 and females.55 In an approach to evaluate human variants of unknown significance in essential meiosis genes, mutations in human MLH1/3 were selected by probing databases such as dbSNP and GnomAD and identifying potential deleterious candidates to introduce into the mouse genome via Crispr/Cas9 editing. Although the fertility of women harboring these mutations is not known, some of the mouse models exhibited premature age-­related infertility and are worth inspecting further.56 After DNA repair is complete, the SC dissolves in the diplonema and diakinesis stages. Little is known about this step of prophase and how it impacts female fertility, but studies in Drosophila

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development

193

9

A Leptonema

DSB formation

Spo11 Endonucleolytic processing Exonuclease processing

Zygonema

Homology searching Single-end invasion

Joint molecule formation Pachynema

B

SDSA

Diplonema

D

Dissolution

F

Nuclease processing

Second-end capture

Strand ligation

Noncrossover

DSBR

C

Resolution As ic y etr pro mme mm ssing y ce S ce ssi tric ng pro E

Strand ligation

Crossover Noncrossover Noncrossover

Crossover

Noncrossover

Crossover

Fig. 9.5 Summary of meiotic recombination based on data from Saccharomyces cerevisiae.  (A) Meiotic recombination is initiated by the formation of DNA double-­strand breaks (DSBs) by the protein Spo11 (orange ellipse) and other accessory proteins. Following DNA cleavage downstream of the DSB event, Spo11 is liberated, forming a Spo11-­oligonucleotide complex. Exonuclease activity generates single-­stranded DNA (ssDNA) that, through the action of the RecA homologs Rad51 and Dmc1, coat the ssDNA, invade homologous DNA templates on the homolog, and create a displacement (D)-­loop. (B) In the following extension of the invading DNA strand, the strand can become disrupted and displaced, reanneal with the opposite side of the DSB, and be repaired as a noncrossover event in a process known as synthesis-­dependent strand annealing (SDSA). Alternatively, the invading strand can continue to become extended creating a larger D-­loop. (C) Upon second-­end capture of the other side of the DSB and subsequent DNA synthesis, a double Holliday junction (dHJ) is formed. (D) By the actions of a helicase and a topoisomerase, dHJs can be migrated toward each other and dissolved (in a process called dissolution) into a noncrossover event. (E) Alternatively, dHJs can be resolved in a symmetrical manner creating noncrossover events or asymmetrically creating crossover events. (F) Following strand invasion, structure-­specific nucleases can process joint molecule intermediates to generate a crossover. Many of these events are drawn largely from studies in S. cerevisiae and can be extrapolated to many meiotic species. Abbreviation: DSBR, double-­strand break repair. (From Gray S, Cohen PE. Control of meiotic crossovers: from double-­strand break formation to designation. Annu Rev Genet. 2016;50:175–210. https://doi.org/10.1146/annurev-genet-120215-035111.)

melanogaster suggest that premature removal of the SC does not affect meiotic chromosome segregation.57 In male mice, removal of the SC requires aurora kinase B.58 In contrast, SCF3 (SKP1– Cullin–F-­box)-­mediated ubiquitin ligase activities maintain the SC while mouse spermatocytes gain competence to complete meiosis.59

OOCYTE GROWTH PHASE O •  ocytes arrest at prophase I for an extended period. • Follicle activation initiates oocyte growth. • Only fully grown oocytes are competent to complete meiosis upon

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PART I  The Fundamentals of Reproduction

A Before luteinizing hormone

B After luteinizing hormone ?

LHR Gs PKA

CNP

EREG

AC

NPR2 MMP

EGFR

EREG

LH LHR Gs

7P

MMP

PKA

High cGMP

CNP NPR2

AC EGFR

Low cGMP P

PDE5

PDE1

PDE5

CNP diffuses extracellularly to cumulus cells

PDE1

CNP and EGFR agonists diffuse extracellularly to cumulus cells

cGMP decreases

High cGMP

cGMP diffuses through gap junctions into the oocyte

GPR3 PDE3A Gs High cAMP AC

Mural granulosa cells

PDE3A

cGMP diffuses through gap junctions out of the oocyte

Meiosis arrested

Cumulus Oocyte cells Gap junction Zona plaques pellucida

Lower activity

GPR3

cAMP Gs decreases AC Meiosis resumes

Higher activity

Fig. 9.6 Working model of signaling pathways that regulate meiotic arrest and resumption in preovulatory follicles. Panel (A) shows a follicle and an expanded view of a mural granulosa cell before LH exposure. Panel (B) depicts events occurring in response to LH. Some of the slower events are not shown, such as the decrease in CNP and the increase in mRNA encoding epiregulin and amphiregulin. Higher levels of enzymatic activity are depicted by darker shades of orange. Abbreviations: AC, adenylyl cyclase; CNP, C-­type natriuretic peptide; EGFR, epidermal growth factor receptor; EREG, epiregulin (and amphiregulin); GPR3, G-­ protein receptor 3; Gs, Gs G protein; LH, luteinizing hormone; LHR, luteinizing hormone receptor; MMP, matrix metalloprotease; NPR2, natriuretic peptide receptor 2; PDE1, 3A, 5, phosphodiesterases; PKA, protein kinase A. (From Jaffe LA, Egbert JR. Regulation of mammalian oocyte meiosis by intercellular communication within the ovarian follicle. Annu Rev Physiol. 2017;79:237–260. https://doi.org/10.1146/annurev-physiol-022516-034102.)

The mammalian oocytes that complete DSB repair and are not subjected to apoptosis make up the ovarian reserve. These oocytes remain arrested in diakinesis until the luteinizing hormone (LH) ovulatory cue triggers the resumption of MI. During the prophase I arrest, oocytes within primordial follicle complexes become activated and start to grow. While growing, oocytes synthesize and accumulate RNAs and organelles that they will need to complete meiosis and to support fertilization and early embryonic development. This oocyte growth occurs at the same time as follicle development and differentiation and is discussed in detail in Chapter 8. It is important here to highlight the long span of time that exists between the completion of recombination and the initiation of meiotic maturation. In mice, oocytes can remain arrested in diakinesis of prophase I for months and in humans, this arrest can last for decades. Mouse spermatocytes, on the other hand, arrest for ∼6 days before resuming and completing MI.

OOCYTE MEIOTIC MATURATION C •  hromatin compaction status is predictive of meiotic competence. • A cAMP/cGMP-­dependent signaling pathway inhibits the resumption of meiosis I in fully grown oocytes; their reduction triggers meiotic resumption, which relies on increased CDK1-­cyclin B1 activity. • Homologous chromosomes tid cohesion at centromeres is protected from proteolysis.

• P  remature separation of sister chromatids is common in oocytes from women of advanced maternal age. • Spindle assembly checkpoint prevents chromosome segregation mistakes and is weakened with age. • Fertilizable eggs arrest at metaphase II due to high CDK1 activity until fertilization. • Meiotic maturation is accompanied by molecular and cellular changes that optimize successful fertilization and embryo development.

Prophase I Arrest and Meiotic Resumption Once oocytes reach their full size (∼80 μm diameter in mice; 120 μm diameter in humans), the chromatin becomes condensed within the nucleus and transcription is silenced.60–61 When detected by microscopy, this condensed chromatin forms a tight ring around the nucleolus and is referred to as the “surrounded” nucleolar (SN) configuration of chromatin. Evidence supports that these oocytes, rather than the nonsurrounded nucleolus (NSN) containing oocytes, are more competent to complete meiosis and support embryonic development after fertilization.62–63 Condensation of the chromatin is regulated epigenetically through acetylation and methylation of histone residues such as trimethylation of histone H3 lysine 9 (H3K9me3) and histone H4 lysine 20.64 In fully grown oocytes, a meiotic arrest is maintained through high intracellular levels of second messengers cAMP and cGMP (Fig. 9.6).65–68 In mice and humans, cAMP is generated

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development

in oocytes through a constitutively active G-­protein coupled receptor, GPR3, and adenylate cyclase 3 (AC3).69,70 The resumption of meiosis is controlled by alterations in the levels and activities of proteins known as cyclins and cyclin-­dependent kinases (CDK), which function as a heterodimer. cAMP elicits a cell-­cycle inhibitor signaling cascade through activation of cyclic AMP-­dependent protein kinase (PKA), which activates WEE2 (WEE2 oocyte meiosis inhibiting kinase) and MYT1 (myelin transcription factor 1) kinases.71 These kinases then phosphorylate and inhibit cyclin-­dependent kinase 1 (CDK1), thereby preventing the breakdown of the nuclear envelope and meiotic resumption. PKA also phosphorylates and inhibits the cell division cycle 25B (CDC25B) phosphatase, an activator of CDK1.72 To keep cAMP levels high in the oocyte, cumulus cells produce cGMP, which reaches oocytes through gap junctions and represses phosphodiesterase 3 (PDE3) so that cAMP cannot be metabolized. Because of the inability to metabolize cAMP, mice lacking the Pde3 gene are infertile because they ovulate prophase I-­arrested oocytes.73 To resume meiosis, cAMP levels must drop. In mature ovarian follicles, the midcycle luteinizing hormone (LH) surge initiates this drop by stimulation of Gs-­ protein-­ coupled LH receptors on follicle cells, resulting in the activation of Gs and generation of cAMP by follicle cell transmembrane adenylyl cyclases. This cAMP subsequently activates transcription factors such as cAMP-­response element binding protein 1 (CREB1) and cAMP-­response element modulator (CREM) that induce or repress the transcription of specific genes important in modulating follicular cell function during oocyte maturation and ovulation. LH-­induced Gs activation also causes dephosphorylation of granulosa cell guanylyl cyclase NPR2, with a consequent reduction in its enzymatic activity.74,75 The resulting dramatic reduction in cGMP levels in the granulosa cells allows rapid diffusion of cGMP out of the oocyte across gap junctions, causing a precipitous drop in oocyte cGMP.76 Lower oocyte cGMP levels are no longer sufficient to inactivate PDE3A, so this phosphodiesterase becomes active and breaks down cAMP, resulting in a loss of the PKA activity required to restrain MPF from inducing meiotic maturation. Studies of meiotic resumption in mouse knockout models have been critical to building our knowledge in this area. For example, in Gpr3 knockout mice, oocytes cannot generate cAMP and therefore fail to maintain the prophase I arrest and spontaneously resume meiosis.77 This resumption occurs because PKA no longer activates WEE2/MYT1 nor does it inhibit CDC25B. Crucially, the LH surge causes a drop in cGMP and cAMP, which alleviates the PKA-­mediated inhibitory signals of CDK1. The inhibitory phosphate placed on CDK1 by WEE2/MYT1 is removed by CDC25B. In mice, removal of the Cdc25b gene causes sterility because of the inability of oocytes to activate CDK1 and resume meiosis.78 CDK1 activity requires binding to cyclin B1. When activated, the complex triggers nuclear envelope breakdown, a morphological marker of meiosis I resumption (Fig. 9.7). As a secondary measure to ensure prophase I arrest, the availability of cyclin B1 is tightly controlled. During the arrest period, this cyclin is subject to proteolysis by the CDH1 (FZR1)-­activated anaphase-­promoting complex/cyclosome (APC/C) and is therefore maintained at low levels. Overexpression of cyclin B1 in mouse oocytes can trigger meiotic resumption,79 and when Cdh1 has been depleted, mouse oocytes undergo precocious meiotic resumption.80 CDH1 activity is therefore also regulated. It is activated by the cell division cycle 14B (CDC14B) phosphatase; depletion of Cdc14b causes precocious meiotic resumption.81 Once activated, cyclin B1 levels stabilize and become sufficient to activate CDK1 to promote exit from the prophase arrest and entry into the meiotic M-phase.

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A

B

C Fig. 9.7 Oocytes at the three most commonly observed stages of meiotic maturation.  (A) Oocyte at the germinal vesicle (GV) stage; note the nucleus with its characteristic single nucleolus (arrow) and granular cytoplasm. (B) Oocyte at metaphase I; note the absence of both a nucleus and a polar body and less granular cytoplasm compared with the GV-­stage oocyte. (C) Egg at metaphase II; note the presence of the first polar body (arrow) and the smooth cytoplasm.

Completion of Meiosis I Completion of MI involves the segregation of homologous chromosomes (Fig. 9.8). The entry into meiotic metaphase (prometaphase I) is a lengthy process (6–7 h versus 1–2 h in mitotic prometaphase) that involves spindle building and resolution of the condensed chromatin. In mice, microtubule organizing centers (MTOCs) are the acentriolar structures that nucleate microtubules to form the MI spindle. Prior to nuclear envelope breakdown, MTOCs first stretch and then fragment.82 This process occurs in at least three phases: (1) restructuring of the MTOCs through aurora kinase A (AURKA) and polo-­like kinase 1 (PLK1) activities, (2) dynein-­ dependent stretching into a ribbon-­ like structure along the nuclear envelope, and (3) fragmentation by kinesin motor protein KIF11 after the nuclear envelope breaks down. After fragmentation and restructuring, the MTOCs sort and cluster into two poles while nucleating microtubules that push and pull resolved chromosomes to the forming metaphase I plate. Mysteriously, a role for MTOCs in building a meiotic spindle in human oocytes may not exist.83 Although some studies describe the presence of MTOCs in discarded human oocytes,84 another study did not detect pericentriolar material and instead demonstrate that microtubules nucleate via a RAS-­ related nuclear protein-­dependent pathway that initiates from chromatin.83 In mice, AURKA and PLK1 are essential for fertility—oocyte-­specific knockouts are sterile—and are required for MTOC fragmentation.85,86 Because of the paucity of human oocyte studies, a comprehensive study of MTOC markers has not yet been conducted. Early prometaphase I steps (i.e., condensing chromosomes, nucleating microtubules, and building a bipolar spindle) require less CDK1 activity. As CDK1 activity increases, later prometaphase I events occur. For example, some microtubules that nucleate from MTOCs will make end-on attachments to kinetochores,

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A

MITOSIS

Cohesin

APCCdc20

SAC * Centromere

APCCdc20

Separase

Securin degraded

Securin

Securin

* Kinetochore Sister chromatids

Prometaphase

Spindle

Anaphase

Metaphase Sister kinetochore biorientation

B

MEIOSIS

APCCdc20

Bivalent

APCCdc20

Chiasma Separase

Cohesin

Securin

Kinetochore Centromere

Spindle

Prophase

Metaphase I Sister kinetochore coorientation

Anaphase I Protection of centromeric cohesin Metaphase II

Homolog biorientation

Anaphase II

Sister kinetochore biorientation

Fig. 9.8 Schematic of chromosome segregation during mitosis and meiosis.  The key adaptations to meiotic chromosomes are indicated. Note that while a single microtubule contacts each kinetochore (mitosis, meiosis II) or pair of kinetochores (meiosis I) in S. cerevisiae, in most organisms, multiple microtubules connect to each kinetochore, resulting in increased probability and configurations of incorrect attachments in both mitosis and meiosis. (A) Key features of mitotic chromosome segregation. (B) Key features and adaptations of meiotic chromosome segregation. (Modified from Marston AL, Wassmann K. Multiple duties for spindle assembly checkpoint kinases in meiosis. Front Cell Dev Biol. 2017;5:109.)

the multiprotein complexes that dock at the centromeres of chromosomes. Other high-­CDK1 activity events include regulating chromosome alignment at the metaphase I plate and migration of the spindle from the center of the oocyte to the cortex.87 Stable kinetochore-­microtubule attachments are essential for triggering anaphase I onset. However, making these attachments is error-­prone; if anaphase were to occur before proper attachments are made, chromosomes can mis-­segregate and result in aneuploidy. Cells employ a spindle assembly checkpoint (SAC) to integrate sensing erroneous attachments with preventing anaphase onset (Fig. 9.9).88,89 The SAC senses kinetochores that are not attached to microtubules. When these kinetochores are sensed, the TTK protein kinase (TTK), more commonly referred to in the literature as MPS1, initiates a signaling cascade to recruit the mitotic checkpoint complex (MCC) to kinetochores.90 In mice, oocytes engineered to express a kinetochore-­binding domain mutant of TTK are subfertile and produce a high percentage of aneuploid eggs.91 In oocytes, the SAC is active early in prometaphase I because no stable kinetochore attachments are yet made.92 The core of the MCC consists of MAD2L1/ BUBR1B/BUB3 (mitotic arrest deficient 2 like 1 / BUB1 mitotic checkpoint serine/threonine kinase B / BUB3 mitotic checkpoint

protein) which binds and sequesters the APC/C activator CDC20 (cell division cycle 20). Without APC/C activity, cyclin B1 levels remain high, and separase remains inactive because it is bound to securin protein. When an inappropriate attachment is made, it is sensed by aurora kinases B and C (AURKB/C) and microtubule depolymerization is triggered.93 This depolymerization thereby creates an unoccupied kinetochore and keeps the SAC activated. Once all attachments are correct and stable, the SAC is satisfied, and CDC20 is freed from the MCC to activate the APC/C. The APC/C targets cyclin B1 and securin for degradation so that metaphase exit and cohesin destruction by separase, respectively, can occur. The result of SAC satisfaction in MI is the separation of homologs where one set remains in the oocyte and the other set is removed in the polar body. Intriguingly, the SAC appears to be less stringent in sensing inappropriate attachments in MI oocytes relative to mitotic cells.94,95 In Mlh1 knockout mice, there are no crossovers and therefore no bivalents. Yet, occasionally oocytes will complete MI and extrude a polar body.96 In contrast, all Mlh1-­null spermatocytes arrest in metaphase, indicating that the stringency of the SAC in male germ cells is more robust than in females.51,52 Other mouse models support the notion that oocytes have a weak SAC response such as XO mice where there is no X chromosome

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development

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Fig. 9.9 Signaling pathway of the spindle assembly checkpoint (SAC).  (A) At anaphase I, correct and full attachment of kinetochores to spindle microtubules leads to disassembly of the mitotic checkpoint complex (MCC) from kinetochores (SAC off). Cdc20 is free to activate the anaphase-­promoting complex/ cyclosome (APC/C), then securin and cyclin B are targeted for degradation. Separase is activated and cleaves the phosphorylated cohesion subunit Rec8 from chromosome arms allowing homologous chromosomes to separate and metaphase I-­anaphase I transition. (B) Unattached kinetochores activate SAC (SAC on). SAC proteins localize to unattached kinetochores and form MCC and sequester Cdc20 preventing it from activation of APC/C. Securin and cyclin B1 levels remain high, separase is not activated, therefore cohesion is not removed and metaphase I-­anaphase I transition is delayed. (Modified from Mikwar M, MacFarlane AJ, Marchetti F. Mechanisms of oocyte aneuploidy associated with advanced maternal age. Mutat Res Rev Mutat Res. 2020;785:108320. https://doi.org/10.1016/j.mrrev.2020.108320.)

homolog and Sycp3 knockout mice that lack crossovers.97–99 Reduced expression of SAC proteins occurs with advanced maternal age and weakening of the SAC is a potential cause of the increase in egg aneuploidy that occurs as women age.100,101 During MI, sister chromatids must remain associated with one another. As women age, premature separation of sister chromatids (PSSC) is the leading defect that drives egg aneuploidy and failure to conceive.102,103 Sister chromatids remain associated through the protection of cohesin complexes at centromeres. This protection occurs through shugoshin proteins (SGO2) that prevent separase cleavage of the guarded cohesin population.104 This cohesion also promotes sister chromatid fusion of kinetochores so that they make attachments to microtubules nucleating from the same spindle pole. This fusion is critical to segregating homologs and not sister chromatids in MI; however, sister kinetochore fusion in human oocytes is surprisingly faulty.105 This lack of kinetochore fusion could explain why errors in female meiosis are common even at young reproductive ages. Cohesin complexes are deposited during premeiotic DNA replication, an event that happens during fetal development in oocytes. As cohesins start to decay over time, they apparently are not replaced.106 107 Therefore, another event driving PSSC with age is cohesin loss,

which can cause sister kinetochore separation and favor attachment of sister kinetochores to microtubules nucleating from opposite poles.108–111 In fact, sister chromatid segregation in MI, also called reverse segregation, occurs frequently in oocytes of older women.102 These separated sister chromatids are at risk for random segregation during meiosis II (MII).

Meiosis II After segregation of homologous chromosomes and an asymmetric cell division with emission of the first polar body, CDK1 activity accumulates rapidly, and oocytes align chromosomes and reassemble a metaphase II spindle. The transition from MI to MII involves skipping DNA replication so that the genome content can become haploid upon completion of MII. Increasing CDK1 activity allows chromosome condensation and kinetochore-­ microtubule connections to occur. Unlike spermatocytes, which will complete MII, the mature oocyte, or preferably, the egg,112 arrests at the metaphase of MII. The metaphase II arrest must be maintained for several hours until fertilization and requires CDK1 activity to remain ). To keep CDK1 activity high, cyclin B1 levels

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remain high by inhibition of the APC/C, as is achieved during the prophase I arrest. However, during metaphase II, the APC/C is inhibited by FBXO43 (previously known as EMI2), thereby allowing CDK1 activity to persist.113–116 Depletion of FBXO43 causes egg activation and metaphase II exit. Upon fertilization, FBXO43-­ mediated inhibition is alleviated by CaMKII (calcium/calmodulin-­dependent protein kinase II) and PLK1 phosphorylation of FBXO43. These phosphorylation events trigger FBXO43 turnover and allow activation of the APC/C to then degrade cyclin B1.114,117 Another protein kinase, called MOS, is also responsible for the arrest of the oocyte cell cycle at metaphase of meiosis II.118–119 Mice deficient in MOS are subfertile because their oocytes do not arrest at metaphase II.120 Instead, the oocytes continue to divide without being fertilized (parthenogenetic activation) and as a result, the mice develop teratomas. With sufficient MOS production, eggs remain arrested in metaphase II until fertilization. The fertilizing sperm induces calcium

CAMKII PLK FBXO43 c-Mos MEK MAPK

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Fig. 9.10 Model of the regulation of MII arrest in mammalian eggs.  High MPF activity is essential for MII arrest and may be maintained via separate pathways; direct inhibition of the APC/C, and direct stabilization of MPF. The pathway that involves EMI2-­mediated CSF arrest is shown in solid lines. In mouse eggs, the c-­Mos pathway is not mediated by p90rsk, so its downstream targets remain obscure (dashed lines), but potential target points are shown as either inhibition of the APC/C or inhibition of EMI2 degradation. MPF activity may negatively regulate the c-­Mos pathway, based on studies on frogs. (Modified from Madgwick S, Jones KT. How eggs arrest at metaphase II: MPF stabilisation plus APC/C inhibition equals Cytostatic Factor. Cell Div. 2007;2:4. https://doi.org/10.1186/1747-1028-2-4.) GV oocyte

Mitochondria

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oscillations that cause cyclin destruction and degradation of MOS protein, resulting in resumption of meiosis II and extrusion of the second polar body. Finally, to separate the sister chromatids, centromeric cohesin must also be removed by separase. During the metaphase II arrest, centromeric cohesin is still protected by SGO2, which also is bound to the PP2A-­B56 phosphatase. For anaphase II onset to occur, an inhibitor of PP2A, called IPP2A, is recruited to allow for deprotection of cohesin and for sister chromatids to segregate.121

Oocyte Maturation-­Associated Changes In addition to meiosis, there are numerous essential molecular and cellular changes that occur during oocyte maturation. These changes include organelle movements, alterations to chromatin epigenetic marks, uptake of essential ions, mRNA degradation, and synthesis and posttranslational modification of essential proteins. The net result of these events is to optimize the ability of the metaphase II-­arrested egg to support successful fertilization and embryo development. At the ultrastructural level, there are changes in the distribution of the organelles, with movement of the endoplasmic reticulum (ER), mitochondria, and cortical granules (CGs) toward the oocyte cortex (Fig. 9.11). Mitochondria become intimately associated with ER tubules including occasional large (1-­2 μm) aggregates of ER tubules that form in cortical locations.122,123 CGs, which continue to be synthesized from Golgi complexes during maturation, move to the cortex where they become anchored just below a subplasma membrane concentration of actin microfilaments. This CG movement occurs along cytoplasmic actin filaments and by “hitchhiking” onto Rab11a-­ positive vesicles that also move to the membrane along actin filaments.124,125 CG anchoring is dependent on the presence of MATER, a protein that serves as an important component of the subcortical maternal complex126 (see Chapter 8). Human cortical granules, unlike in the mouse, are found around the entire periphery of the oocyte, even in regions immediately adjacent to the MI and MII spindles.123,127 By the time of metaphase II arrest, Golgi complexes have largely disappeared, explaining a decline in the ability of the mature egg to synthesize new secreted proteins. In addition, the transzonal projections responsible for cumulus cell-­ oocyte coupling have retracted by MII, disrupting gap junctional communication between the oocyte and cumulus cells.128 With movement of chromatin to the cortical region, the oocyte becomes highly asymmetric (Fig. 9.11). The actin cytoskeleton is altered, with thickening of cortical actin overlying the metaphase II spindle. This region of the plasma membrane is devoid of microvilli—unlike the rest of the oocyte plasma membrane, which is enriched with microvilli. This loss of microvilli may reduce the chance of sperm entering the region of the

MII egg

Fig. 9.11 Schematic representation of subcellular morphological patterns in germinal vesicle (GV) oocytes, metaphase I (MI) oocytes, and metaphase II (MII) eggs. (Modified from Trebichalská Z, Kyjovská D, Kloudová S, Otevřel P, Hampl A, Holubcová Z. Cytoplasmic maturation in human oocytes: an 2021;104(1):106–116.

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development

metaphase II spindle where it could interfere with the normal progression of meiosis. The spindle is maintained in the cortical region by actin-­dependent cytoplasmic streaming driven by an actin nucleator, the ARP2/3 complex.129 Concomitant with chromosomal condensation and movements orchestrated by the spindle during meiotic maturation, the oocyte chromatin continues to undergo epigenetic remodeling that was initiated during oocyte growth (see Chapter 8). More extensive domains of the noncanonical broad histone H3 lysine 27 trimethyl (H3K27me3) marks are detected in eggs than in full-­grown oocytes.130 This finding indicates that there is continued activity during maturation of lysine methyltransferase 2B, the major enzyme responsible for establishing these marks.130,131 It is also likely that lysine-­specific demethylase 4A continues to remove H3K9me3 marks, preventing disruption of the broad H3K27me3 domains.132 Finally, DNA methylation at imprinting control regions of some genes is not complete until following oocyte maturation.133 Zinc is an essential transition metal that serves both structural and catalytic roles within numerous cellular proteins such as zinc-­ finger transcription factors and metalloenzymes. Extensive zinc accumulation via uptake through plasma membrane zinc transporters occurs during oocyte maturation, with an overall increase in zinc content of about 50%.134,135 The reduction of available zinc in prophase I-­arrested oocytes causes resumption of meiosis by perturbing the function of the zinc-­binding protein FBXO43 and also causes abnormalities in cortical reorganization leading to diminished cell polarity.136,137 Zinc is stored in the cortical granules in preparation for its release during fertilization.138 Calcium is also taken up during oocyte maturation and stored in the endoplasmic reticulum. Calcium enters the oocyte’s cytoplasm through at least two distinct calcium channels: the voltage-­ gated T-­ type channel, CaV3.2, and the constitutively active transient receptor potential channel, TRPM7.139,140 Cytoplasmic calcium is then actively transported into the ER through sarcoendoplasmic reticulum calcium ATPase (SERCA) pumps. Calcium influx during oocyte maturation is essential for appropriate calcium signaling to occur at fertilization. Oocyte maturation is accompanied by the recruitment of specific dormant maternal mRNAs for translation into protein. This recruitment provides a mechanism for the transcriptionally silent oocyte to initiate a new developmental program required for fertilization and embryo development. Dormant maternal mRNAs have in common that they encode proteins that are critical for functions in the oocyte during maturation, in the egg at fertilization, or in the very early stages of embryo development. In rodents, examples of these recruited mRNAs are Mos, the inositol 1,4,5-­triphosphate receptor, type 1 (Itpr1), and tankyrase 1. As mentioned above, translation of Mos is critical for blocking cell cycle progression beyond metaphase II. The maturation-­ associated increase in ITPR1 protein is important for successful egg activation at fertilization by increasing the ability of the egg to exhibit calcium oscillations.141 Tankyrase 1 has a critical role in regulating ß-­catenin transcriptional activity in the zygote and 2-­cell embryo, which is essential for development beyond the 2-­cell stage.142 The molecular mechanism by which dormant maternal mRNAs are recruited is cytoplasmic polyadenylation. Stored maternal mRNAs are released from subcortical aggregates in response to CDK1-­ mediated phosphorylation of the RNA-­ binding protein MSY2 (Y box-­binding protein 2) at the initiation of oocyte maturation.143 The dormant maternal mRNAs have specific nucleotide sequences in the 3′ untranslated region called cytoplasmic polyadenylation elements (CPEs). CPEs direct the binding of poly(A) polymerase and the addition of poly(A) tracts to dormant maternal mRNAs. Among the proteins that regulate this process are cytoplasmic polyadenylation binding element protein 1 (CPEB1) and deleted in azoospermia-

199

(DAZL), which itself is a dormant maternal mRNA.144,145 Polyadenylation leads to association with polysomes, translation of the mRNAs, and consequently to increases in the levels of the encoded proteins. Most maternal mRNAs present in the oocyte are not recruited for translation but instead undergo rapid degradation in preparation for a switch from the maternal to the embryonic transcriptional program. These mRNAs lack CPEs and do not undergo polyadenylation and translation to generate proteins. Instead, they are targeted by different enzymes responsible for promoting mRNA decay. For example, maternal mRNAs are destabilized by deadenylation, which is mediated by the CCR4-­NOT deadenylation complex, prior to degradation by the exosome complex.146,147 Terminal uridylyl transferases 4 and 7 (TUT4 and TUT7) place a uridine onto the 3′ terminus of mRNAs that have a short polyA tail.148,149 The 3′ uridylation promotes entry into both the 5′-­to-­3′ and 3′-­to-­5′ degradation pathways. The decapping enzymes DCP1A and DCP2 promote mRNA degradation by removing the 5′ monomethyl guanosine cap in preparation for mRNA entry into the 5′-­to-­3′ mRNA degradation pathway.150 Of note, mRNAs encoding DCP1A, DCP2, and two components of the CCR4-­ NOT deadenylation complex (CNOT6L and CNOT7) are all dormant maternal mRNAs recruited for translation during oocyte maturation, consistent with their critical roles in maturation-­associated RNA degradation. In human embryos, the CCR4-­NOT deadenylation complex appears to participate in maternal mRNA degradation.151 Low expression of transcripts encoding CCR4-­NOT proteins is found in embryos that fail to cleave to the 2-­cell stage. These findings indicate that as in the mice, maternal mRNA degradation during oocyte maturation is essential for successful development. Synthesis and posttranslational modification of cytoplasmic proteins also occur during oocyte maturation. For example, glutathione is synthesized during maturation from amino acid precursors; a sufficient level of this reducing agent is required for initial events of fertilization.152 Microtubules undergo posttranslational alterations in acetylation during the transition from metaphase I to metaphase II.153 Microtubule acetylation is essential for proper organelle positioning and movement, possibly by affecting interactions between tubulin and cytoplasmic lattices.154 Phosphorylation and dephosphorylation of cytoplasmic proteins, particularly those involved in regulating the cell cycle, are required for successful cytoplasmic maturation. Together, resumption of meiosis until arrest at metaphase II and extensive changes to chromatin, the contents of the cytoplasm, and organelle positioning combine to generate a mature egg that is poised for successful fertilization and embryo development. Defects in any of these processes can lead to failed fertilization, suboptimal embryo development, and even failed implantation and, consequently, impaired fertility.

SPERM TRANSPORT • S  perm transport from the vagina to the site of fertilization (ampulla of the Fallopian tube) is promoted by uterine contractions. • Sperm stored in the female reproductive tract for up to six days can result in pregnancy. • Sperm transit toward the ampulla involves a series of specific sperm binding and release interactions with the female reproductive tract epithelium. • Sperm may be assisted in directional targeting to the ampulla by fluid flows (rheotaxis), chemical signals (chemotaxis), and temperature changes (thermotaxis).    Sperm have a significant challenge to overcome following ejaculation: they need to travel a long distance (relative to the size of the sperm) to the site of fertilization within the female reproductive tract. This process is highly inefficient because a very low

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percentage of ejaculated sperm ever reach the site of fertilization, the ampulla of the Fallopian tube (human oviduct). The idea that sperm transit is a mechanism for stringent selection of only the most fecund sperm is supported by animal studies but has not been shown in humans.155–157

Female Reproductive Tract Contributions to Sperm Transport Sperm are rescued from the acidic vaginal environment by the presence of cervical mucus secretions generated during the periovulatory time period. Sperm can survive and retain their motility in endocervical crypts for up to 5 days, and the cervical mucus there provides energy substrates important for sperm survival, including glucose and fructose.158,159 However, some sperm transit very rapidly from the cervix to the Fallopian tube, with documentation of transit times as low as 5 minutes following insemination.160 Sperm motility is not required for this transit because inert beads or protein aggregates placed in the vaginal fornix can be found in the uterus and Fallopian tubes.161,162 Uterine contractions may promote sperm migration toward the Fallopian tubes, accounting in part for this rapid transit time. This is true in mice, in which live imaging experiments show an essential role of uterine contractility in sperm entry into the oviduct.163 In humans, contractile waves from the uterine cervix to the fundus occur in the late follicular phase.164 These uterine contractions may be promoted following coitus by factors in semen such as prostaglandins. The number of uterine contractions following intrauterine insemination correlates with live birth, consistent with the idea that uterine contractions may assist sperm transit.165 However, it is unknown if the sperm that rapidly transit to the Fallopian tubes with assistance from uterine contractions are actually capable of fertilizing eggs. Instead, the sperm that arrive later may be responsible for achieving pregnancy.

Sperm Interactions With the Female Reproductive Tract The observation that, in humans, sexual intercourse six days before ovulation can result in pregnancy suggests that there is long-­term storage of functional sperm in the female reproductive tract.166 The uterine environment can be detrimental to sperm viability, so mechanisms are in place to protect sperm from uterus-­derived secreted cytotoxic factors. In the mouse, seminal vesicle secretory protein 2 (SVS2) is a highly abundant semen component that binds to sperm in the uterus and is required for long-­term sperm survival in the female reproductive tract.167 In humans, sperm are stored in endocervical crypts, from which motile sperm can be retrieved several days after intercourse. Although sperm release from endocervical crypts over time could explain pregnancy long after intercourse, as yet there is no evidence supporting this idea. In a number of animal species, the isthmus region of the oviduct serves as a storage reservoir for sperm, but such a region has not been identified in human Fallopian tubes.168 Sperm interact with the epithelium of the female reproductive tract through specific sperm cell surface molecules. Bovine sperm interact with the epithelium through BSP (binder of sperm) proteins, which are heparin-­binding acidic proteins made in seminal vesicles and adsorbed onto the sperm surface following ejaculation.169,170 Mouse sperm lacking cell surface ADAM3 (a disintegrin and metallopeptidase domain 3, also known as cyritestin) are unable to migrate through the uterotubal junction, presumably because they fail to bind the oviductal epithelium.171 ADAM proteins are cell surface proteases that also interact with integrins, explaining their cell adhesion function, though an oviductal epithelial cell surface binding partner for ADAM3 has not been identified.

The oviductal epithelial surface molecules that interact with the sperm have been identified in the cow to be annexins. These proteins are present on bovine oviductal epithelial apical membranes and bind to purified BSPs; antibodies to annexins impair sperm binding to the epithelium.172 Although there is no evidence for annexins as epithelial sperm receptors in humans, two annexin proteins, ANXA1 and ANXA5, are highly abundant in the Fallopian tube in the early secretory phase and could serve in this function.173 Based on studies in several species, sperm-­epithelium binding is likely mediated by carbohydrate molecules on the epithelial proteins.170 In cows and pigs, the relevant carbohydrate is fucose; BSP1 was identified based on its fucose-­binding ability and annexins contain fucose. Sialic acid appears to be important for sperm-­epithelium interactions in the hamster and horse. Gradual sperm transit toward the ovulated egg appears to involve a series of sperm binding interactions with the epithelium. These binding events help to maintain sperm viability and motility, likely through inhibiting sperm maturation changes and oxidative membrane damage.174–177 Sperm capacitation (see Sperm capacitation below) is associated with decreased epithelial binding and hyperactivated sperm motility, a vigorous whip-­ like pattern that can help sperm detach from the epithelium and ascend toward the egg.178,179 Sperm lacking the ability to hyperactivate do not progress beyond the oviductal isthmus because they fail to detach.180 Therefore, capacitation-­associated changes permit sperm to progress up the reproductive tract by a series of binding and release events. The extensive mucosal folds present in the uterus and Fallopian tubes create physical barriers for the sperm to overcome on their journey and may select against less motile sperm, though in some regions they create channels to direct the sperm toward the egg.

Cues That Direct Sperm Migration In  vitro, sperm are assisted in directional movements by fluid flows (rheotaxis), chemical signals (chemotaxis), and temperature changes (thermotaxis), but it is not clear how much these environmental cues contribute to sperm transit in vivo.181–183 Sperm swim upstream against slow fluid flow, so fluid coming from the direction of the ovulated egg would promote sperm migration in that direction. However, a recent study demonstrated that fluid flows within the mouse oviduct go in the opposite direction; from the uterus toward the ovary.184 Such flows could promote sperm movement toward the egg but suggest that rheotaxis is not physiologically significant, at least in the mouse. Chemotaxis and thermotaxis impact motility by altering the beat pattern of the sperm tail, the flagellum. Chemotaxis is thought to influence the directional movement of sperm that are close to the ovulated egg rather than regulating long-­distance movements. For example, a local progesterone gradient generated by cumulus cells changes the motility characteristics of nearby sperm from progressive to hyperactivated.185,186 There is even evidence that human sperm have different chemotactic responses to follicular fluid in different women, indicating that chemotaxis could influence fertility.187,188 Sperm can sense changes in temperature along a gradient and tend to move in the direction of the warmer temperatures present at the oviductal ampulla relative to the isthmus.189,190 Together, these physical and chemical cues may promote the success of sperm migration to reach the ovulated egg.

SPERM CAPACITATION • S  perm undergo capacitation (develop the capacity to fertilize an egg) during transit through the female reproductive tract. • Capacitation entails numerous biochemical and functional changes including alterations in sperm membranes, motility patterns, ion transport, and signaling pathways.

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development

• C  alcium plays a central role in the regulation of capacitation-­ associated changes in progressive sperm motility, hyperactivation, and acrosomal exocytosis. • CatSper is the principal calcium channel in sperm and must be functional for the development of hyperactivated motility. • Development of the capability to undergo acrosomal exocytosis is a key aspect of sperm capacitation.    Like oocytes, sperm must undergo a maturation process before they can participate in a successful fertilization event. The observation that following ejaculation, sperm must spend several hours in the female reproductive tract before being able to fertilize ovulated eggs in vivo was made independently by Austin and Chang in the early 1950s.191,192 Austin later coined the term capacitation for this sperm maturation process because it entails development of the capacity to fertilize an egg.193 A chemically defined culture medium that supports sperm capacitation in vitro was later reported by Yanagimachi and Chang.194 This finding led eventually to the widespread ability of scientists to perform in vitro fertilization without first incubating sperm in the female reproductive tract. Many functional and biochemical changes necessary for capacitation have been elucidated. Generally speaking, capacitation includes alterations in sperm membranes, motility patterns, ion transport, and signaling pathways. Key events of sperm capacitation are the development of hyperactivated motility and development of the capacity to undergo acrosomal exocytosis, a membrane fusion event that results in the release of the acrosomal contents. Much of the research on capacitation has been done using animal models, but the ready availability of human sperm for study has made it possible to identify molecular changes that are both similar to other animals and unique to human sperm.

Sperm Membrane Changes Sperm membranes change early on in the process of capacitation. The membranes of ejaculated sperm are coated with seminal fluid that contains “decapacitation factors,” which can reverse the process of capacitation, and other factors that simply inhibit capacitation.195 Several such factors have been identified in animals, including glycoproteins, polysaccharides, and membrane-­ bound vesicles that can serve as cholesterol donors.196–201 In humans, semenogelin, a major component of semen, immobilizes ejaculated sperm and prevents other capacitation-­associated changes.202,203 Cholesterol in human seminal plasma also inhibits capacitation.204 Sperm lose these factors as they progress through the female reproductive tract or when incubated in culture media in vitro, allowing capacitation to begin. Capacitation is associated with an increase in sperm membrane fluidity,205 which is initiated by exposure of sperm to the relatively high levels of bicarbonate in the female reproductive tract. Bicarbonate and calcium influx stimulate the activity of soluble adenylyl cyclase, a sperm tail enzyme that generates cAMP and promotes PKA activity.206–208 PKA subsequently triggers a cascade of protein phosphorylation events thought to be important for the success of capacitation.209–211 One response to this bicarbonate-­activated phosphorylation cascade is that the sperm plasma membranes, which before capacitation have a high degree of phospholipid asymmetry in the inner and outer leaflets, undergo “scrambling” or redistribution of the phospholipids.212 Phospholipid redistribution, along with loss of membrane cholesterol due to efflux onto cholesterol acceptors, reduces membrane stability and allows for diffusion of sperm membrane proteins into regions that support appropriate sperm-­egg interactions.213 The increase in membrane fluidity may also promote fusibility of the sperm membranes. Media used for capacitating sperm in vitro contain proteins such as albumin, which serves as a cholesterol acceptor, promoting the removal of cholesterol from the sperm membranes.

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Sperm Ionic Changes Human sperm capacitating in vitro gradually undergo alkalinization, which appears to require cholesterol efflux.214 Alkalinization stimulates membrane hyperpolarization, calcium influx, and sperm motility. The increase in intracellular pH is mediated by various ion channels and transporters, including proton and bicarbonate transporters and the cystic fibrosis transmembrane conductance regulator (CFTR), which is a chloride-­ selective anion channel that can also transport bicarbonate.215,216 In mouse sperm, there is a sperm-­specific sodium-­proton exchanger (encoded by Slc9a10) that mediates alkalinization due to proton extrusion.217 Knockout mice lacking this exchanger have impaired motility and are completely infertile. In human sperm, alkalinization is mediated by hydrogen voltage-­ gated channel 1 (HVCN1).215 Interestingly, HVCN1 is potently inhibited by zinc, so the zinc-­ associated decrease in sperm motility following fertilization (see Blocks to polyspermy below) could be mediated by HVCN1. CFTR function is regulated by phosphorylation by PKA, so activation of the cAMP/PKA signaling pathway during capacitation provides a mechanism for controlling CFTR function and sperm intracellular pH.218 CFTR also interacts with other bicarbonate transporters that regulate its function. Male mice heterozygous for a null mutation in Cftr do not undergo capacitation normally— they have impaired sperm alkalinization, reduced cAMP levels, and reduced fertility both in vitro and in vivo.219 These findings, combined with studies of human sperm using chemical inhibitors of CFTR, suggest that men heterozygous for Cftr mutations may have infertility due to abnormal capacitation.220 Similarly, mutations in SLC26A3, a chloride-­ bicarbonate exchanger that modulates CFTR function, are associated with male subfertility.221 In addition to sperm becoming alkalinized, their membranes become hyperpolarized, i.e., have a reduction in membrane potential, during capacitation.222 Hyperpolarization of human sperm requires PKA activation and is necessary for appropriate patterns of sperm motility through its impact on intracellular calcium levels and transport of other ions. Sperm membrane potential is regulated in part by the actions of two potassium channels, SLO1 and SLO3.223–225 In addition, proton efflux channels, sodium/proton exchangers, sodium/bicarbonate cotransporters, epithelial sodium channels, and sodium/potassium pumps, such as the testis-­specific Na,K-­ATPase alpha 4 isoforms, contribute to the regulation of membrane potential.215,226,227 Calcium plays a central role in the regulation of capacitation and sperm function. It has key roles in sperm motility, hyperactivation, and acrosomal exocytosis. Sperm intracellular calcium levels, which increase during capacitation, are regulated by the balance of activities of calcium influx channels and efflux pumps combined with sequestration into and release from intracellular stores. The main calcium stores in sperm are the acrosome, a large Golgi-­derived vesicle overlying the nucleus, and the redundant nuclear envelope in the neck region.228–230 Many of the molecular mediators of calcium transport in sperm have been identified using chemical inhibitors, knockout mouse models, and documentation of mutations in men with abnormalities in sperm function. As in somatic cells, calcium release from intracellular stores appears to be largely controlled by the activity of the IP3 receptor (IP3R), but the ryanodine receptor may also play a role. SERCA pumps and secretory pathway calcium ATPase pumps use ATP to transport calcium against a concentration gradient into sperm calcium stores. Sperm calcium efflux to the external environment is controlled by both plasma membrane calcium ATPase pumps (PMCA) and sodium/ calcium exchangers.231,232 Many calcium influx channels are present in sperm. Of these, channels identified to have critical functions include the 2.3 channel, which regulates sperm motility

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CatSper Ca2+

Longitudinal column of the fibrous sheath

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Flagellar cytoplasm EFCAB9

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Fig. 9.12 Structure of the mammalian CatSper channel and its localization in the sperm flagellum.  (A) Located in the principal piece of the sperm flagellum, the heteromeric voltage-­dependent, pH-­sensitive CatSper channel is composed of ten subunits. The channel consists of four pore-­forming α subunits (CatSper 1–4) as well as five auxiliary subunits: CatSperβ (beta), CatSperγ (gamma), CatSperδ (delta), CatSperζ (zeta), and CatSperε (epsilon). A new member of the CatSper complex has been recently identified as EF-­hand calcium-­ binding domain-­containing protein 9 or EFCAB9, a calmodulin-­like protein that binds calcium and acts as a dual calcium and pH sensor for CatSper. (B) CatSper channels form functional domains within the principal piece to enable rapid propagation of the calcium signals along the flagellum. The channel complex is organized in quadrilateral longitudinal nanodomains that form a unique pattern of four linear threads running down the principal piece of the flagellum. (From Rahban R, Nef S. CatSper: The complex main gate of calcium entry in mammalian spermatozoa. Mol Cell Endocrinol. 2020;518:110951. https://doi.org/10.1016/j.mce.2020.110951.)

and acrosomal exocytosis, and the sperm-­specific cation channel of sperm (CatSper), which is required for hyperactivation.233–235 Indeed, human male infertility can be explained by mutations in proteins that comprise or interact with the CatSper channel.236–240 CatSper, the principal calcium channel in sperm, is a complex ion channel comprised of four pore-­forming alpha subunits and six accessory subunits (Fig. 9.12).241 It is localized to the principal piece of the sperm tail in highly organized, longitudinal stripes, with each stripe containing two rows of CatSper complexes positioned on each side of the two longitudinal fibers of the fibrous sheath.242 This positioning is maintained by interactions between one of the CatSper accessory subunits, Catsperζ, and EFCAB9 (EF-­ hand calcium-­ binding domain-­ containing protein 9).240 Positioning along the full length of the principal piece allows for linear propagation of calcium signals along the sperm tail. The bilaterally, but not quadrilaterally symmetrical positioning of the longitudinal stripes appears to be important for regulating the asymmetric motility pattern characteristic of hyperactivation. CatSper is regulated by changes in intracellular pH and by ligands present in oviductal fluid such as prostaglandins and progesterone.243 The channel is constitutively active, but calcium influx through CatSper is potentiated by alkalinization of the sperm cytosol and upregulated in response to cAMP-­dependent PKA activity.244,245 An interesting, nongenomic signaling pathway (see Chapter 5) controls CatSper-­ mediated sperm chemoattraction responses to progesterone released by cumulus cells. CatSper activity is normally restrained by the endocannabinoid 2-­ arachidonoylglycerol, an endogenous eicosanoid derived from arachidonic acid and present in the sperm plasma membrane.246 Progesterone binds to a sperm membrane progesterone receptor, a/b hydrolase domain-­containing protein 2 (ABHD2). In response to progesterone binding, ABHD2 cleaves 2-­ arachidonoylglycerol, relieving inhibition of CatSper and allowing calcium influx. Calcium influx-

hyperactivation responses directs sperm toward the source of progesterone, the cumulus-­oocyte complex.

Competence for Acrosomal Exocytosis Development of the capability to undergo acrosomal exocytosis is a key functional aspect of sperm capacitation. The acrosome contains hydrolytic enzymes that may help the sperm penetrate through the oocyte’s outer vestments—the cumulus cells and zona pellucida (ZP). The inner acrosomal membrane is tethered to the nuclear envelope whereas the outer acrosomal membrane lies adjacent to the plasma membrane. Acrosomal exocytosis occurs when the sperm’s outer acrosomal membrane fuses in multiple locations with the sperm plasma membrane (Fig. 9.13). This process releases the soluble acrosomal contents and exposes the particulate acrosomal matrix and inner acrosomal membrane to the extracellular environment. Acrosomal exocytosis results in exposure of membrane components and proteins that are essential for sperm-­egg interactions. Capacitation-­associated loss of sperm membrane cholesterol, which increases membrane fluidity, may increase the competence of these membranes to fuse. As with most exocytotic events, acrosomal exocytosis is calcium regulated, so the capacitation-­associated increase in intracellular calcium levels increases sperm competence for this event.

FERTILIZATION • F  ertilization occurs in the ampullary region of the Fallopian tube. • Penetration of the hyaluronic acid matrix within the cumulus cell mass is assisted by hyaluronidase enzymes tethered to the sperm head. • Sperm bind to zona pellucida protein ZP2 and penetrate the zona with mechanical assistance from hyperactivated sperm motility; it is not clear if there is essential role for acrosomal enzymes or sperm-­

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Fig. 9.13 Stages of acrosomal exocytosis.  (A) Capacitated, acrosome-­intact human sperm. The acrosome (A) overlies the anterior portion of the nucleus (N). The equatorial segment (ES) is a narrowing of the posterior region of the acrosome. Bar = 0.25 μm. Inset: Membranes of the sperm head. The plasma membrane (PM) is the most external and the outer acrosomal membrane (OAM) lies directly beneath it. The inner acrosomal membrane (IAM) lies on the inner surface of the acrosome, and the nuclear envelope (NE) separates the nucleus from the acrosome. Bar = 0.10 μm. (B) Swelling of the acrosomal matrix anterior to the equatorial segment. Bar = 0.16 μm. (C) The plasma membrane and outer acrosomal membrane begin to fuse in the anterior region, indicated by arrowheads and the area of vesiculation (V). Bar = 0.20 μm. (D) Membrane fusion complete, excluding the equatorial segment. Vesicles separate from sperm head. Bar = 0.19 μm. (E) Vesicles separate from anterior sperm head, leaving the inner acrosomal membrane as the new outer boundary of anterior sperm. The plasma membrane of the equatorial segment remains. Bar 0.16 μm. (Modified from Yudin AI, Gottlieb W, Meizel S. Ultrastructural studies of the early events of the human sperm acrosome reaction as initiated by human follicular fluid. Gamete Res. 1988;20(1):11–24. https://doi.org/10.1002/mrd.1120200103.)

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Fig. 9.14 Steps involved in sperm approach and fusion with egg.  (A) Acrosomal exocytosis occurs at some point before sperm penetrate the zona pellucida, either in the oviduct, within the hyaluronic acid matrix of the cumulus cell layer, or upon reaching the zona pellucida. During acrosomal exocytosis, IZUMO1 on the outer acrosomal membrane migrates out to sperm plasma membrane at the equatorial segment (red). Acrosome contents shown in green. (B) Zona pellucida penetration. (C) Acrosome-­reacted sperm bind at the equatorial segment to the egg plasma membrane through an IZUMO1-­IZUMO1R (JUNO) interaction and then fuse with the egg. CD9 on the egg surface is essential for sperm-­egg fusion. (Modified from Okabe M. Mechanisms of fertilization elucidated by gene-­manipulated animals. Asian J Androl. 2015;17(4):646–652. https://doi.org/ 10.4103/1008-682X.153299.)

• S  perm-­egg plasma membrane adhesion is mediated by the sperm head protein IZUMO1 and the egg IZUMO1 receptor, IZUMO1R. • Sperm-­egg plasma membrane fusion is orchestrated by several proteins on both sperm and egg, but no specific fusogenic protein has been identified. • Following fusion, the egg actively incorporates the sperm into its cytoplasm and the sperm chromatin decondenses, allowing protamine-­ histone exchange.    Fertilization, or the union of sperm and egg, occurs in the ampullary region of the Fallopian tube. The ovulated egg is surrounded by an expanded mass of cumulus cells and the ZP, both of which the sperm needs to penetrate before reaching and fusing with the egg plasma membrane. The cumulus mass, ZP, and egg plasma membrane each provide an opportunity for the selection of only the most fecund sperm by ensuring that it has all the necessary enzymes and binding partners to successfully negotiate these barriers. Following fertilization, changes to the ZP and egg plasma membrane prevent polyspermy, or fertilization by additional sperm, which precludes normal embryo development.

Sperm Penetration of Egg Vestments During the process of meiotic maturation, the cumulus cells secrete hyaluronic acid, which along with additional secreted proteins and components from the serum becomes incorporated into a hyaluronan extracellular matrix separating the cumulus cells from each other (see Chapter 8). Sperm penetrate this matrix with the assistance of a hyaluronidase, SPAM1, that is tethered to the sperm plasma membrane by a glycosyl phosphatidylinositol anchor.247

Even after acrosomal exocytosis, SPAM1 remains associated with the newly exposed inner acrosomal membranes, facilitating penetration of the cumulus mass by acrosome-­reacted sperm (Fig. 9.14). Mouse and rat sperm have a second plasma membrane-­associated hyaluronidase, HYAL5, but the gene encoding this protein is not present in humans.248 The presence of HYAL5 likely explains the ability of mouse sperm lacking SPAM1 to penetrate the cumulus mass, though they do so more slowly than wild-­type sperm. Penetration of the cumulus layer brings the sperm to the next barrier—the egg’s extracellular coat, the ZP. The human ZP is comprised of four glycoproteins: ZP1, ZP2, ZP3, and ZP4 (previously ZPB) (see Chapter 8).249 Its structure is very similar to the ZP of rodents except that rodents have only three ZP proteins. Initial studies in mice concluded that sperm bind to specific sugar moieties, N-­or O-­glycans, that are attached to ZP3. However, more recent work using knockout mouse models in which the individual mouse ZP proteins were replaced by their human ZP protein counterparts have shown conclusively that this is not the case.250,251 Instead, human sperm bind to the N-­terminal region of human ZP2, and there is no requirement for sugar moieties for this interaction.138,252 Several candidate sperm proteins capable of binding to the ZP have been identified, including β-­1,4-­ galactosyltransferase, milk fat globule-­EGF factor 8 (previously SED1), zona pellucida 3 receptor (previously sp56), and zonadhesin, but the corresponding knockout mice were fertile.253–256 These knockout experiments indicate that the individual proteins are not essential, but do not exclude the possibility that more than one of these sperm molecules could participate in sperm-­ZP binding. The identity of the molecule (or molecules) on human sperm that interact with egg ZP2 is unknown but could be an excellent target for contraceptive development.

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development Fig. 9.15 Fertilization requires two distinct sperm-­egg fusion steps.  (A) Acrosome-­intact sperm. (B) Acrosomal exocytosis causes IZUMO1 translocation to the equatorial segment plasma membrane where it is accessible for binding to egg IZUMO1R. Some IZUMO1 remains associated with the inner acrosomal membrane at the anterior sperm head. (C) IZUMO1-­IZUMO1R interactions, combined with interactions between additional sperm and egg proteins, leads to the first sperm-­egg fusion event. Sperm-­egg fusion activates cortical actin-­mediated cytoplasmic protrusions. (D) Engulfment of the sperm head by cytoplasmic protrusions leads to a second fusion event that resembles endocytosis. Green, acrosomal contents; red circles, IZUMO1; blue circles, IZUMO1R; purple circles, CD9. (Modified from Okabe M. The cell biology of mammalian fertilization. Development. 2013;140(22):4471–4479. https://doi.org/10.1242/dev.090613.)

How sperm penetrate the ZP is not entirely clear, although a hyperactivated motility pattern is essential and acrosomal exocytosis must occur at some point prior to complete ZP penetration, because sperm recovered from the perivitelline space, the region between the ZP and egg plasma membrane, are always acrosome-­ reacted.257–260 The classical model explaining ZP penetration was that sperm binding to ZP3 at the ZP surface induces acrosomal exocytosis, releasing enzymes responsible for local dissolution of the ZP, allowing sperm to penetrate. In mice, not only do sperm bind ZP2 rather than ZP3, but sperm can undergo acrosomal exocytosis in the oviduct prior to reaching either the ZP or the cumulus cells surrounding the egg.261–263 Furthermore, sperm that previously underwent acrosomal exocytosis remain capable of penetrating the ZP.260,264 These findings demonstrate that soluble acrosomal enzymes are not required for ZP penetration, though they could still facilitate the process. For example, acrosin and PRSS21, two serine proteases located in the acrosome, are not absolutely required for fertilization in the mouse.265,266 However, mouse sperm lacking either one of these proteases do not fertilize eggs in vitro as quickly as do wild-­type sperm, and double knockout mice lacking both proteases are subfertile in vivo.266–268 In addition, hamster sperm lacking acrosin cannot fertilize eggs in vivo, even though they reach and bind to the ZP of ovulated eggs in the ampulla.269 These findings indicate that acrosin has an essential role in ZP penetration in hamsters and perhaps also in other species. The only sperm protein identified to date as being essential for ZP penetration is the sperm membrane protein ADAM3 (a disintegrin and metallopeptidase domain 3, previously known as cyritestin).270 The mechanistic role of ADAM3 in ZP penetration is unclear, though it could participate in ZP binding and/or facilitate ZP penetration through its protease activity. Additional proteins identified in mouse knockout studies to be required for ZP penetration all impact the expression of ADAM3 on sperm. These proteins include calmegin, a testis-­specific chaperone protein required for proper ADAM protein folding; two other ADAM proteins, ADAM1A and ADAM2, without which ADAM3 is not expressed; and angiotensin I converting enzyme, which impacts distribution of ADAM3 on the sperm surface.271–274 Interestingly, lack of these same proteins is associated with a failure of sperm to migrate through the uterotubal junction, supporting a role for ADAM3 in that process as well.171

Sperm-­Egg Adhesion and Fusion The final barrier to fertilization is the egg plasma membrane. Sperm adhere to the egg through interactions at the equatorial segment, the region of the plasma membrane of the sperm head immediately adjacent to the exposed inner acrosomal membrane (Fig. 9.15).275 Following adhesion, the sperm and egg undergo a plasma membrane fusion process that brings the sperm con tents into continuity with the egg cytoplasm. Gamete adhesion

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and gamete fusion appear to be mechanistically distinct processes because the gametes can bind to each other successfully and yet not undergo fusion. Several protein mediators of gamete fusion have been identified, but despite decades of research, a mechanistic understanding of gamete fusion is lacking. Sperm-­egg plasma membrane adhesion is mediated by sperm IZUMO1, which was named after a Japanese shrine dedicated to marriage, and the egg IZUMO1 receptor, IZUMO1R (also known as JUNO, the Roman goddess of fertility and marriage, and previously known as folate receptor 4, FOLR4).258,276 IZUMO1 is an immunoglobulin superfamily, type I membrane protein that has an N-­terminal “Izumo domain,” a single immunoglobulin-­like domain, a single transmembrane domain, and a short C-­terminal intracellular tail. IZUMO1 is localized to the acrosomal membranes, but during acrosomal exocytosis, it becomes accessible to the extracellular milieu when the inner acrosomal membrane becomes exposed and when IZUMO1 translocates from the outer acrosomal membrane to the equatorial segment of the plasma membrane277 (Fig. 9.15). The N-­terminal Izumo domain is responsible for binding to IZUMO1R.278 IZUMO1R is localized to the external surface of the entire egg plasma membrane.276 It has a globular structure and is attached to the egg membrane by a glycosyl phosphatidylinositol anchor.276,279,280 Structural analysis of human IZUMO1 with IZUMO1R demonstrates a high-­affinity interaction that causes a conformational change in IZUMO1 and results in the formation of a stable complex.279,280 Unlike sperm-­egg adhesion, sperm-­egg fusion requires multiple distinct proteins. On the sperm side, SPACA6 (sperm acrosome-­ associated protein 6), SOF1 (spermoocyte fusion required 1), FIMP (fertilization-­ influencing membrane protein), and TMEM95 (transmembrane protein 95) are required for sperm-­ egg fusion but not adhesion.281–284 TMEM95 was first identified in a genome-­wide association study of bulls whose sperm had exceptionally poor performance when used for artificial insemination.285 These bulls were homozygous for a nonsense mutation in TMEM95 that was later shown to impair the ability of their sperm to fuse with bovine eggs.286 Similarly, mouse sperm lacking TMEM95 penetrate the ZP and bind to the plasma membrane of mouse eggs but fail to fuse.281,284 TMEM95 is lost from bull sperm during acrosomal exocytosis, suggesting it has an indirect role in fusion by facilitating fusogenic functions of other proteins. FIMP and SOF1 were identified as testis-­enriched genes in the mouse, but they are widely conserved in other mammals including humans.281,282 There are two isoforms of FIMP, one with a transmembrane domain and one that is secreted. Mouse sperm lacking both FIMP isoforms can bind but not fuse to eggs, but only the transmembrane isoform is necessary for fusion activity. Mouse sperm lacking SOF1 have essentially the same phenotype—they are capable of penetrating the ZP and binding the egg plasma membrane but fail to fuse, leading to complete

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Spaca6 was first associated with infertility in transgenic male mice generated during a large-­scale mutagenesis screen to search for genes involved in reproduction.283 SPACA6 is an immunoglobulin superfamily protein with a single transmembrane domain that has structural homology to IZUMO1. Also, like IZUMO1, it localizes to the acrosome and translocates to the equatorial segment following acrosomal exocytosis.281 Mouse sperm lacking SPACA6 penetrate the ZP and adhere to the egg plasma membrane but fail to fuse, phenocopying FIMP, SOF1, and TMEM95. The identification of three distinct proteins (FIMP, SOF1, and SPACA6), all of which are required to function within the sperm plasma membrane to support fusion events suggests that these proteins participate in a multiprotein complex that orchestrates sperm-­egg fusion. IZUMO1 may also participate in sperm-­egg fusion through such complexes. IZUMO1 is associated with multiprotein complexes identified in mice, rat, and hamster sperm using biochemical techniques.287 Participation of IZUMO1 in these multiprotein complexes requires the C-­terminal cytoplasmic tail and/or transmembrane domains, whereas the N-­terminal Izumo domain only mediates homodimer formation. These protein-­protein interactions could lead to the formation of large protein aggregates in the sperm membranes. In fact, IZUMO1 interacts with FIMP, SOF1, and SPACA6 when the proteins are overexpressed in cultured somatic cells.281 However, whether or not these interactions occur between the endogenous proteins present in sperm is not known. Further studies are needed to identify the full composition of these multiprotein complexes and test their functional roles in sperm-­egg fusion. A major outstanding question is the identity of a sperm protein with characteristics of a membrane fusogen that could mediate sperm-­egg fusion. Two egg plasma membrane proteins, CD9 and CD81, are important for sperm-­egg fusion. They are members of the tetraspanin superfamily, a group of four-­transmembrane domain proteins that typically serve as scaffolds for macromolecular complexes that function in activities including cell–cell adhesion and signal transduction. Mouse eggs lacking CD9 support sperm binding but not fusion, leading to severe infertility.288–290 The role of CD81 is not as critical as that of CD9 because the fertility of female mice lacking CD81 is only reduced by 40%.291 However, mice lacking both CD9 and CD81 are completely infertile, suggesting that they have similar functions in sperm-­ egg fusion. In fact, sperm-­egg fusion in eggs lacking CD9 can be partially rescued by overexpressing CD81, indicating that CD81 can compensate for the function of CD9.292 It is not clear exactly what CD9 and CD81 do to support fusion, but the most compelling model is that they together recruit additional egg proteins into a macromolecular complex that carries out signaling or other activities necessary for membrane fusion. One such activity is likely to be supporting the normal structure and dynamic state of egg microvilli. Within microvilli, CD9 interacts with two proteins that link it to the actin core.293 Mouse eggs lacking CD9 have very abnormal short, thick microvilli that do not appear to be dynamic. The thick microvilli have an increased radius of curvature, which would make membrane fusion more challenging. A recruited signaling molecule that could participate in fusion is integrin α6β1, which interacts with mouse egg CD9 based on a coimmunoprecipitation assay.289 Integrins are heterodimeric proteins that link the extracellular matrix to the cortical cytoskeleton but can also mediate cell–cell adhesion and transmit signals that change cell behavior. In the mouse, inhibition of integrin α6β1 using either function-­blocking antibodies or peptides interferes with sperm-­egg fusion, but knockout studies indicate that neither integrin α6β1 nor any other β1-­containing integrin is essential for this process.288,289 294–296 As with sperm, no egg protein with

Fig. 9.16 Scanning EM of human sperm being incorporated into hamster egg.  Egg cytoplasmic processes extend over the equatorial segment. (From Tsuiki A, Hoshiai H, Takahashi K, Suzuki M, Hoshi K. Sperm-­egg interactions observed by scanning electron microscopy. Arch Androl. 1986;16(1):35–47. https://doi.org/10.3109/01485018608986921.)

characteristics of a membrane fusogen has yet been discovered to serve a functional role in sperm-­egg fusion.

Sperm Incorporation Following the initial fusion of the egg and sperm plasma membranes, the sperm becomes incorporated into the egg cytoplasm, and the sperm nucleus decondenses. These events are largely orchestrated by the egg, whereas the sperm is a passive participant. Sperm attached to the egg surface are quickly engaged by egg microvilli, which surround and trap the sperm head, particularly in the region of the equatorial segment and postacrosomal head region297–300 (Fig. 9.16). Shortly before membrane fusion, the sperm flagellum stops beating.301 With the advent of sperm-­ egg fusion, the egg cortex becomes “activated” to generate cytoplasmic protrusions that extend over and eventually completely cover the sperm head. This process results in the formation of a “fertilization cone,” an actin-­rich structure that temporarily overlies the newly incorporated sperm head. The egg cortical movements are actin-­mediated and are regulated by Rho family GTPases.302–-­305 Once the sperm head enters the egg cytoplasm, the sperm nuclear envelope disappears first in the region toward the base of the sperm head. Subsequently, the inner acrosomal membrane peels away from the anterior region of the sperm nucleus and the remainder of the sperm nuclear envelope disappears.277,299 Sperm nuclear decondensation begins when the nucleus becomes exposed to the egg cytoplasm. Sperm protamines become oxidized during epididymal maturation, creating protamine disulfide bonds that stabilize the nuclear structure. These disulfide bonds are reduced by egg cytoplasmic glutathione, a major intracellular source of free thiol groups, opening up the densely packed sperm chromatin.152,306 When this happens, protamines are rapidly replaced by maternal histone proteins, allowing the sperm chromatin to regain a nucleosome-­ based structure. Phosphorylation of sperm protamines contributes

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Fig. 9.17 Cycle of Ca2+ transient generation in mammalian eggs at fertilization.  Starting at the top left, the large grey arrows show temporal order. For each panel, the cytoplasmic Ca2+ trace is colored orange at the portions of the trace that are generated mainly due to the steps illustrated in that panel. Top left: Sperm PLCζ acts on PIP2 in intracellular vesicles to generate IP3, which stimulates IP3R-­mediated Ca2+ release and subsequent Ca2+-­induced Ca2+ release. Top right: Ca2+ stimulates mitochondrial ATP production; ATP is required for SERCA pump activity. Bottom right: Ca2+ is pumped back into the ER through SERCA pumps and out of the egg through PMCA pumps and NCX. Bottom left: Ca2+ flows into the cytoplasm through TRMP7, CaV3.2, and TRPV3 channels and is then available for SERCA pumps to replenish ER Ca2+ stores in preparation for the next Ca2+ release event. Orange dots indicate Ca2+ at its destination; small grey arrows show the direction of flow. CaV3.2, T-­type voltage-­dependent Ca2+ channel; IP3, inositol trisphosphate; IP3R, IP3 receptor; MCU, mitochondrial uniporter; NCX, sodium/Ca2+ exchanger; PIP2, phosphatidylinositol 4,5-­bisphosphate; PLCζ, phospholipase C zeta; PMCA, plasma membrane Ca2+ ATPase; SERCA, sarco/ endoplasmic reticulum Ca2+ ATPase pump; TRPM7, transient receptor potential cation channel subfamily M member 7; TRPV3, transient receptor potential cation channel subfamily V member 3. (Modified from Stein P, Savy V, Williams AM, Williams CJ. Modulators of calcium signalling at fertilization. Open Biol.

2020;10(7):200118.)

to protamine-­ histone exchange by facilitating interactions between protamines and the highly abundant egg acidic protein, nucleoplasmin.307–309

EGG ACTIVATION • E  gg activation causes the resumption of meiosis II and second polar body emission and converts the fused sperm and egg into a totipotent zygote. • A sperm-­specific phospholipase C, PLCζ, is released into the egg upon sperm-­egg fusion and initiates calcium release from endoplasmic reticulum stores. • Continued oscillations in intracellular calcium promote egg activation events; the major downstream mediator is calcium/calmodulin-­ dependent protein kinase II. • The calcium oscillatory pattern at fertilization, which is regulated by calcium pumps and influx channels, drives the success of development and offspring health. • Several blocks to polyspermy are established during egg activation that ensure fertilization by a single sperm. • The male and female pronuclei form in the periphery of the zygote and then replicate their DNA while migrating centrally in preparation for the first mitotic division.    Egg activation is a term that encompasses the numerous events required to convert the metaphase II-­arrested egg that has just fused with sperm into a totipotent zygote, or 1-­cell embryo. These events include calcium oscillations, cortical granule exo cytosis, resumption of meiosis with emission of the second polar

body, pronucleus formation, initiation of chromatin reprogramming, and establishment of the blocks to polyspermy. Much of the literature regarding the mechanisms mediating egg activation is based on studies in rodents or domestic animals. However, more recent studies of human preimplantation embryos have demonstrated many similarities; a few differences will be highlighted.

Induction and Regulation of Calcium Oscillations The initial event of egg activation is a prolonged (several minutes) rise in egg cytoplasmic calcium levels due to release from ER calcium stores.310 Calcium release is initiated when a sperm-­specific isoform of phospholipase C, PLCζ, diffuses from the sperm head into the egg cytoplasm following sperm-­egg fusion.311 Although it was initially identified in the mouse, the importance of PLCζ for egg activation in humans was definitively demonstrated with the identification of an infertile man with mutations in both PLCZ1 alleles.312,313 These mutations led to fertilization failure following intracytoplasmic sperm injection during clinically assisted reproduction. A number of additional PLCZ1 mutations associated with male infertility have since been identified.314–317 PLCζ cleaves a membrane-­associated phospholipid, phosphatidylinositol 4,5-­bisphosphate, generating the second messenger, inositol 1,4,5-­trisphosphate (IP3) (Fig. 9.17). IP3 binds to IP3 receptors, which are extremely large protein complexes that form calcium channels in ER membranes. IP3 receptors open in response to the binding of IP3 and calcium, resulting in the rapid release of calcium into the cytoplasm.318,319 This calcium release, combined with persistent generation of IP3 by PLCζ, provides

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continued stimulation of additional IP3 receptor opening events in nearby locations, eventually leading to a wave of cytoplasmic calcium that propagates across the egg. When the ER calcium stores become depleted and cytoplasmic calcium levels rise sufficiently to inhibit IP3 receptor opening, no further calcium is released. Although eggs are resilient enough to handle large increases in cytoplasmic calcium, they eventually return their cytoplasmic calcium levels to baseline through the actions of ATP-­dependent calcium pumps. The ATP required for these pumps is generated by mitochondria in response to the calcium released from the ER.320,321 Calcium is pumped back into the ER by SERCA pumps; this activity helps replenish ER calcium stores. Calcium is also pumped outside of the egg through the actions of plasma membrane calcium ATPases (PMCA). PMCA actions reduce the calcium content of the egg, so to fully replenish ER calcium stores, calcium must be allowed back into the egg through plasma membrane calcium influx channels. In the mouse, there are at least three channels that support calcium influx following fertilization: the T-­type voltage-­gated channel, CaV3.2, and two different transient receptor potential channels, TRPM7 and TRPV3.140,322,323 Disrupting the function of these channels results in a failure to replenish ER calcium stores. TRPM7 is highly sensitive to the amount of calcium and magnesium in culture media, and alterations in the concentrations of these ions affect how well TRPM7 supports calcium influx and replenishment of ER calcium. A special feature of calcium signaling at fertilization in mammals, including humans, is that rather than a single increase in calcium as in most other animals, there is a repetitive series of oscillations in cytoplasmic calcium following fertilization that lasts for several hours.310 These oscillations occur because of the continued presence of sperm PLCζ generating IP3. When ER calcium stores are replenished due to the combined actions of calcium influx channels and SERCA pumps, IP3 and cytoplasmic calcium binding to IP3 receptors can once again cause calcium release into the cytoplasm, starting a new cycle of calcium release and reuptake. Calcium oscillations last variable amounts of time due to differences in the amount of sperm PLCζ released into the egg, the level of calcium in the ER, the amount of ATP generated to power the PMCA and SERCA pumps, and desensitization of the IP3 receptor to ligand stimulation. There is strong evidence in mouse models that the calcium oscillatory pattern at fertilization drives the success of egg activation, preimplantation embryo development, implantation, development to term, and offspring growth.324–326 These findings were first demonstrated by studies in which an electroporation apparatus was used to entrain different patterns of calcium oscillations on recently fertilized zygotes. The zygotes were then transferred to pseudopregnant dams and evaluated for the above parameters. A decreased level of calcium exposure led to a reduction in implantation efficiency and impaired development to term. An increased level of calcium exposure did not impact implantation but reduced development to term and increased weight variation in offspring. There was no impact of altered calcium signaling on the morphological appearance of the blastocyst stage embryos flushed from the dam, but there were significant gene expression differences in the blastocysts that likely explained the abnormal implantation and development. These studies used a very artificial system to entrain calcium oscillations, but similar observations have been made using knockout mouse models that have reduced calcium signals at fertilization in vivo.140,322 Together, these studies highlight the critical importance of the calcium oscillatory pattern for the success of development.

Calcium-­Induced Signaling One of the most important targets of calcium signaling in the egg is CaMKII. CaMKII proteins are serine/threonine kinases

that exist in cells as an assembly of multiple oligomers arranged in a ring shape.327 Each oligomer has an autoinhibitory domain that keeps the protein inactive by preventing access of substrates to the catalytic domain. Rising intracellular calcium levels cause calcium binding to calmodulin, a small regulatory protein that amplifies cellular calcium signals. Calcium/calmodulin binding to CaMKII causes a conformational change that moves the autoinhibitory domain away from the catalytic region of the protein. In response, CaMKII becomes active and autophosphorylates itself, preventing rebinding of the autoinhibitory domain, and also phosphorylates substrate proteins. CaMKII becomes inactive only when it is dephosphorylated. In this way, CaMKII translates information encoded in calcium signaling into phosphorylation events that alter downstream protein functions or stability. In the fertilized egg, CaMKII activity increases and decreases in synchrony with calcium oscillations, so a continued pattern of calcium oscillations causes persistent intermittent spikes in CaMKII activity.324 The critical nature of CaMKII activity for egg activation was first demonstrated by the observation that overexpression in mouse eggs of a constitutively active form of the CaMKII alpha isoform could activate the eggs sufficiently to support full-­term development, even when calcium oscillations were completely suppressed.324 The identity of the CaMKII isoform responsible for mediating egg activation was discovered when mouse eggs completely lacking the gamma isoform, CaMKIIγ, had normal calcium oscillations at fertilization but failed to resume meiosis.328 This failure is most likely because, in the absence of CaMKIIγ, FBXO43 is not phosphorylated and destroyed by the ubiquitin/proteasome system, so meiosis resumption is prevented (see Meiosis II section above). A second calcium-­dependent response of egg activation is CG exocytosis. Within a few minutes of the first calcium rise, subplasmalemmal actin is cleared in localized regions, allowing trafficking of the CGs to the plasma membrane. Actin clearing and CG trafficking require both the presence of MATER and the activity of the nonmuscle actin motor protein, myosin IIA.126 Similar to vesicular exocytosis in somatic secretory cells, CG exocytosis in eggs is mediated by soluble N-­ethylmaleimide-­sensitive factor attachment protein receptor (SNARE) proteins. In this pathway, an increase in calcium causes SNAREs present on both the target membrane (t-­SNAREs) and the vesicle membrane (v-­SNAREs) to form a complex with each other. Additional proteins stabilize the SNARE complex, causing the two membranes to come into close contact and fuse, releasing the vesicle contents. Based on studies in mouse and porcine eggs, SNARE pathway proteins involved in CG exocytosis include the t-­SNARE, SNAP23, the v-­SNARE, VAMP1, Rab family GTPases, soluble NSF attachment protein, and N-­ethylmaleimide sensitive factor.329–332

Blocks to Polyspermy CG exocytosis is a critical event of egg activation because it helps to prevent polyspermy, or entry of more than one sperm into the egg, which is incompatible with normal development. CG exocytosis has been known for decades to cause “hardening” of the ZP as indicated by a resistance to proteolytic or heat-­mediated digestion. Physical changes to the ZP following fertilization include stiffening and thinning of the ZP and condensation and fusion of ZP filament bundles.333–336 Biochemical changes are also observed, including cleavage of ZP2 into two fragments that remain connected by disulfide bonds and an increase in the amount of zinc in the ZP.334,337 Together, these changes to the ZP not only prevent additional sperm from fertilizing eggs, forming the “ZP block to polyspermy,” but also have a role in protecting the newly formed embryo in transit through the female reproductive tract. Ovastacin, a zinc metalloendopeptidase, was the first CG component to be definitively identified as participating in the ZP block to polyspermy. When released from the egg during

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1. Zona penetration ~minutes post fertilization

Zona pellucida Periviteline space

Zinc release

2. Membrane block ~minutes post fertilization IZUMO1 JUNO (vesicles) Fertilized egg

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methods including chemical methods of increasing intracellular calcium, overexpression of CaMKII, or intracytoplasmic sperm injection. Of note, although it is possible that CG exocytosis contributes to the membrane block, additional mechanisms are required for its establishment. One likely mechanism is shedding from the egg surface of membrane-­bound vesicles that contain sperm binding molecules including IZUMO1R and CD9.276,340 These vesicles can serve as sperm decoys by supporting sperm binding away from the egg plasma membrane, precluding a productive sperm-­egg fusion event. The third type of polyspermy block is rapid but transient. Zinc released from CGs rapidly diffuses through the ZP where it interferes with the motility of supernumerary sperm, likely because of its inhibitory effects on sperm alkalinization and calcium influx.138,215 Sperm with impaired motility do not efficiently penetrate the ZP. However, with continued diffusion, the zinc concentration becomes too low to impact sperm motility and this block is not maintained.

Pronucleus Development and Migration

Oolemma 3. Zona block ~4 hour post fertilization ZP4 ZP3 ZP2 ZP1

*

Ovastacin

Fig. 9.18 Postfertilization blocks to polyspermy.  After fertilization and cortical granule exocytosis, two immediate blocks and one delayed block occur to prevent polyspermic fertilization. (1) Zinc release: zinc released within a few minutes of fertilization affects the forward motility of sperm that are binding or have initiated penetration of the zona. (2) Membrane block: ∼40 minutes after fertilization, IZUMO1R(JUNO) is shed from the oolemma within vesicles (green) into the perivitelline space to act as a decoy that blocks any supernumerary sperm from adhering and fusing with the oolemma. (3) Zona block: within 4 hour postfertilization, ovastacin cleaves ZP2 at the N terminal region (asterisk) and the cleavage of ZP2 provides a definitive block by preventing further sperm binding to the zona pellucida. (Modified from Bhakta HH, Refai FH, Avella MA. The molecular mechanisms mediating mammalian fertilization. Development. 2019;146(15):dev176966. https://doi.org/10.1242/dev.176966.)

CG exocytosis, ovastacin cleaves ZP2 near its N-­terminus, disrupting the sperm binding site and preventing additional sperm from binding and penetrating the ZP338 (Fig. 9.18). ZP2 cleavage is a stable mechanism, but it occurs relatively slowly (over the course of several hours) as calcium oscillations continue to induce waves of CG exocytosis and ovastacin release. There is also evidence that zinc released from CGs and incorporated into the ZP changes ZP architecture in a way that impairs sperm binding.138,334 A second type of block to polyspermy is known as the “membrane block” because it prevents fertilization by sperm that have already penetrated the ZP and have access to the egg plasma membrane. This block is established between 45 minutes and a few hours after fertilization in the mouse as indicated by experiments in which fertilized eggs had their ZP removed and then were inseminated again.339 Little is known about the mechanism(s) by which the membrane block is established, but it requires sperm-­ egg plasma membrane interactions because it does not occur when egg activation is achieved by artificial

Following completion of meiosis II and formation of the fertilization cone, the male and female pronuclei form as nuclear membranes coalesce around the decondensing chromatin. The timing of pronucleus formation coincides with a significant decrease in the activity of mitogen-­activated protein kinase (MAPK).341,342 The pronuclei migrate toward each other at the center of the zygote under the control of two distinct mechanisms. The pronuclei are first pushed away from the zygote cortex due to local nucleation of actin in the region of the fertilization cone (male pronucleus) or oocyte cortex (female pronucleus).343 The second phase of pronuclear migration in the central region of the zygote is mediated by microtubules and the microtubule-­based motor protein, dynein. This phase also depends on the presence of an intact actin cytoskeleton that provides a cortical actin structure that the microtubules push against. In the mouse, the microtubules are nucleated by γ-­tubulin at multiple acentriolar “microtubule organizing centers” that coalesce into spindle poles on opposite sides of the pronuclei. In humans, the microtubules are nucleated by paternal centrioles initially located in the sperm neck region, the junction of the sperm nucleus and flagellum. One of these centrioles, the proximal centriole, for many years was thought to be the only centriole inherited by the human zygote. However, recent studies demonstrated the presence of an atypical distal centriole in sperm that provides a second paternally inherited centriole.344 This centriole requires maternal components to form a complete centrosome. The two centrioles duplicate themselves prior to the first embryonic mitosis, so each daughter cell inherits two centrioles. DNA replication (S-­phase) occurs during pronuclear migration and is followed by a long G2 phase of the cell cycle during which chromatin reprogramming begins (see Embryonic genome activation below). This first mitotic cell cycle takes more than 24 hours, but mitosis itself takes less than 15 minutes to complete.345 Mitosis occurs after the male and female pronuclei reach each other in the center of the zygote. It begins with pronuclear envelope breakdown, followed by chromosome condensation, formation of the mitotic spindle, alignment of the condensed chromosomes along the metaphase plate, chromatid separation, and then symmetrical cleavage to form two similar-­size blastomeres, generating the 2-­cell stage embryo (Fig. 9.19). There is evidence in the mouse that rather than a single spindle, the first mitosis is carried out by two closely aligned but not merged bipolar spindles, one for each pronucleus.346 This mechanism of cell division would explain the observation of the physical separation of the male and female chromatin into different nuclear compartments in the 2-cell stage embryo.347 Whether or not there

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Reprogramming the Embryo Cytoplasm

A

B

C

D

Fig. 9.19 Development of human embryos during the first 72 hours after insemination.  (A) A zygote at 18 hours, showing two polar bodies (arrows) and two pronuclei in apposition (arrowheads) with several distinct nucleoli polarized toward the juxtaposed nuclear membranes. (B) A two-­cell embryo at 28 hours. (C) A four-­cell embryo at 44 hours. (D) An eight-­cell embryo at 68 hours.

is a dual spindle and differential compartments for maternal and paternal chromatin in human 2-­cell stage embryos is not known.

CLEAVAGE STAGE DEVELOPMENT • T  he cleavage stage embryo (2-­ to 8-­cell) undergoes dramatic changes to the cytoplasm and nuclei as the embryo begins functioning as a new individual with a distinct genome. • The embryo recycles materials inherited from both gametes, removing unneeded proteins, mRNAs, paternal mitochondria, and other cellular components. • New embryonic transcripts are generated in 2-­cell human embryos but major embryonic genome activation (EGA) occurs at the 4-­ to 8-­cell stage. • Chromatin reprogramming, or global changes in chromatin composition and organization, occurs during the cleavage stages and is essential for the success of EGA. • Both maternal factors and embryonic transcription factors drive chromatin reprogramming and initiation of EGA.    Cleavage stage development begins at the 2-­cell stage and ends with the transition to the compacted morula stage, when the embryo has 8 to 16 cells. It consists of simple, symmetrical cell division events that generate smaller and smaller blastomeres, with no overall increase in embryo size. These early embryonic cell cycles are slow, lasting from 18 to 24 hours.345 Cleavage stage development is accompanied by dramatic biochemical changes to the cytoplasmic and nuclear contents as the embryo turns over materials inherited from the gametes and begins functioning as a new individual with a distinct genome. These biochemical changes include removal of unneeded parts of the fertilizing sperm, degradation of maternal RNAs and proteins, and the initiation of transcription from the newly formed embryonic genome, or embryonic genome activation. Throughout the cleavage stage, the embryo chromatin undergoes a reprogramming process that is required to change the inherited, fully differentiated sperm and egg chromatin into that of a totipotent embryo.

Most parts of the sperm are engulfed by the egg at fertilization, but the majority of these are degraded during the cleavage stages. For example, the sperm tail initially remains associated with the male pronucleus and can be observed in the 2-­cell stage embryo but is not present after that time.348 Sperm mitochondria are actively targeted for degradation within the egg cytoplasm, likely by at least two mechanisms. Sperm mitochondrial proteins are ubiquitinated during spermatogenesis, identifying them as distinct from maternal mitochondria.349 Ubiquitination marks the paternal mitochondrial proteins for degradation by the proteasome. Ubiquitination can also lead to autophagy, a distinct mechanism for regulated lysosomal degradation and recycling of proteins, organelles, and other subcellular components. Paternal mitochondria are incorporated into autophagosomes through the actions of maternal cytoplasmic ubiquitin ligases and an autophagy adapter protein, sequestosome 1.350,351 The autophagosomes then merge with lysosomes that degrade their contents, allowing the early embryo to recycle amino acids and other macromolecules. It has also been proposed that sperm mitochondrial DNA is degraded within sperm prior to fertilization and that the mitochondria are not degraded but instead are lost over time.352 In any case, targeted destruction and/or lack of paternal mitochondrial replication explain why mammalian mitochondria are maternally inherited.353 Degradation of maternal mRNAs begins during oocyte maturation but continues in cleavage stage embryos through the 8-­to the 10-­cell stage.354 This process is critically important for complete conversion from a maternally directed to an embryo-­ directed developmental program and provides nucleic acids to be recycled into embryonic transcripts. Mediators of maternal mRNA degradation are likely to be similar to those active during oocyte maturation: 3ʹ-­ terminal uridylyl transferases, decapping enzymes, and deadenylases. There is also evidence from studies in the pig that maternal mRNA degradation can be mediated by autophagy.355 Human embryos express mRNAs encoding TUT4 and TUT7 beginning at the 8-­cell stage, and embryos that arrest development at the 8-­cell stage have abnormally low levels of TUT7 mRNA.151 This finding suggests that 3ʹ-­terminal uridylation is an important pathway for mRNA degradation in the human. Degradation of maternal proteins is also essential for the success of cleavage stage development. The autophagy pathway is one major mediator of this process.356 Mouse embryos lacking oocyte-­derived maternal autophagy-­related protein 5 (ATG5), a protein required for autophagosome formation, do not develop beyond the 4-­to 8-­cell stage unless they are rescued by ATG5 from wild-­type sperm.357 These embryos produce 30% less protein than normal, suggesting that a failure to recycle amino acids required for synthesis of new proteins is the cause of their developmental arrest. The ubiquitin-­proteasome pathway is a second major mediator of maternal protein degradation. This pathway depends on a large family of E3 ubiquitin ligases that covalently link ubiquitin molecules to specific proteins that are then directed to the proteasome for degradation. For example, RNF114, an E3 ubiquitin ligase, is required for mouse embryo development because it ubiquitinates TAB1 (TAK-­1 binding protein 1), targeting it for destruction by the proteasome.358 TAB1 must be cleared from the early embryo to allow translocation of nuclear factor kappa B (NF-­kB) from the cytoplasm to the nucleus, which is required for development beyond the 2-­cell stage.358,359 A second example is the ubiquitination of maternally derived axin proteins in early embryos. Proteasomal degradation of axin, a component of the β-­catenin “destruction complex,” prevents phosphorylation and targeted destruction of β-­catenin in zygotes and 2-­cell stage embryos.142 Axin degradation is required to allow β-­catenin-­ mediated transcription necessary for embryo development.

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Metaphase II egg

PN0–PN3

PN3b–PN5

Two-cell

Four-cell

mRNA level

High Maternal mRNA

Minor wave

Eight-cell

Morula

Blastocyst

Major wave

Embryonic mRNA

Low DNA methylation

High 5mC 5mC Low

5hmC Fig. 9.20 Timing of changes in transcription and DNA methylation in human embryos.  Maternal mRNA stores begin to decline during oocyte maturation and continue to decrease through the two-­cell stage (orange line). A minor wave of embryonic mRNA transcription begins during the two-­cell stage and continues until the onset of the major wave between the four-­and eight-­cell stages (green line). Paternal DNA methylation (5mC) is high prior to fertilization but actively removed in the male pronucleus by conversion to 5-­hydroxymethylcytosine (5hmC) (blue solid and dashed lines). Maternal DNA methylation is gradually lost during DNA replication (red line). (Modified from Eckersley-­Maslin MA, Alda-­Catalinas C, Reik W. Dynamics of the epigenetic landscape during the maternal-­to-­ zygotic transition. Nat Rev Mol Cell Biol. 2018;19(7):436–450. https://doi.org/10.1038/s41580-018-0008-z.)

Chromatin Reprogramming and Embryonic Genome Activation Embryonic genome activation (EGA) is one of the most critical cleavage stage events. It traditionally was considered the time when development becomes dependent on protein products of embryonic transcripts. More recently, EGA has been defined as a major increase in transcripts generated from the embryonic genome. The timing of EGA varies in different species, ranging from the 2-­cell stage in the mouse to the 8-­to 16-­cell stage in cows and rhesus monkeys.360–362 In human embryos, some embryonic transcripts are generated as early as the 2-­cell stage, with a gradual increase in transcript numbers through the 4-­cell stage.354 Between the 4-­and 8-­cell stages, there is a major increase in expression of embryonic transcripts, or “major EGA,” and new embryonic proteins can be detected at the 8-­cell stage363,364 (Fig. 9.20). These findings are similar to those in the mouse, in which a “minor EGA” occurs in the zygote prior to major EGA at the 2-­cell stage.365–368 The minor EGA in mice is promiscuous, with many transcripts generated from intergenic regions not controlled by defined core promoters.369 EGA is tightly intertwined with the process of chromatin reprogramming. If EGA is prevented by the use of transcription inhibitors, chromatin reprogramming is impaired, and similarly, inhibition of chromatin reprogramming prevents EGA. Chromatin reprogramming in the cleavage stage embryo involves a resetting of the mature, differentiated chromatin state inherited from the sperm and egg to the largely naïve state required for totipotency. Chromatin is organized in loops of DNA around nucleosomes, which are further organized into dense arrays in regions of “heterochromatin” that effectively restrict interactions with other proteins. Alternatively, in euchromatin, the nucleosomes are not densely packed and can be displaced by the actions of transcription factors and chromatinremodeling proteins. Active gene transcription occurs in regions

of euchromatin. Chromatin reprogramming occurs at multiple levels, including changes in DNA methylation, histone variants included in nucleosomes, histone posttranslational modifications, nucleosome positioning, and chromatin domains. These changes regulate how transcription factors and other transcription machinery can access genomic loci, strongly influencing gene regulation. One of the earliest chromatin reprogramming events is global DNA demethylation, which begins soon after fertilization.370 The paternal genome is >80% methylated in sperm, which is much higher than the maternal genome in eggs (∼55%). Active demethylation of the paternal genome is carried out by TET3, a methylcytosine dioxygenase that converts methyl groups to hydroxymethyl groups, which can then be removed enzymatically to generate a nonmodified cytosine.371,372 DNA repair mechanisms may also contribute to the active demethylation of the paternal genome. The maternal genome is also demethylated after fertilization, but to a much lesser extent, such that in zygotes both genomes are ∼50% methylated. Demethylation continues as preimplantation development progresses, reaching a nadir of ∼25% at the blastocyst stage. Importantly, the methylation of imprinted genes is maintained despite this global demethylation process. Although demethylation predominates, de novo methylation also occurs in specific regions of the genome, particularly during the 4-­to 8-­cell stages when a set of active DNA repeat elements are methylated, suppressing their ability to destabilize the genome. Changes in the specific histone variants incorporated into nucleosomes are another important aspect of chromatin reprogramming. As mentioned above, sperm nuclear decondensation is accompanied by the replacement of protamines with egg histones. In the mouse, the major egg histone H3 variant incorporated into the paternal chromatin is the histone H3.3, which is associated with an open chromatin state.373–375 H3.3 is also incorporated into the maternal chromatin, but at a later pronuclear stage.373

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H3.3 helps to maintain pronuclear chromatin in a decondensed state by preventing the incorporation of histone H1, a linker histone that facilitates chromosome condensation.376 H3.3 incorporation depends on the activity of the histone chaperone, HIRA (histone cell cycle regulation defective homolog A), without which the male pronucleus does not form and the female genome does not support development.377 Histone posttranslational modifications strongly influence chromatin packaging and thus recruitment of transcription factors and gene expression. For example, H3 lysine 27 acetylation (H3K27ac) is an epigenetic mark associated with “open” chromatin comprised of active gene regulatory elements such as promoters and enhancers of genes being transcribed. In mice, H3K27ac increases dramatically following fertilization to reach a peak at the 2-­cell stage; these marks are enriched near genes expressed during EGA.378 Histone H3 lysine 4 trimethylation (H3K4me3) is canonically associated with transcriptional start sites of active genes. However, as mentioned previously (see Chapter 8), full-­grown oocytes have noncanonical broad domains of histone H3K4me3 that are associated with transcriptional quiescence.130,378,379 These broad domains are gradually removed by the time of major EGA due to the actions of histone lysine demethylases, relieving gene repression. In 2-­cell embryos, H3K4me3 is once again mainly restricted to canonical peaks associated with active gene transcription. There is minimal information available regarding histone modifications in human cleavage stage embryos, but immunofluorescence studies show a reduction in the levels of H3K4me3 and H3K27me3 between the 4-­and 8-­cell stages before they increase again in blastocysts.370 These findings suggest that like in the mouse, histone modifications play an important role in chromatin reprogramming in the early human embryo. Chromatin accessibility is a physical indicator of which areas of genomic DNA can be contacted by nuclear macromolecules such as transcription factors. In mouse embryos, chromatin is highly accessible prior to EGA.380,381 This high accessibility is accompanied by low level, highly promiscuous transcription from both protein-­coding loci and intergenic regions that lack promoter elements, including transposons.382 Human zygotes and 2-­cell embryos also have widespread accessible chromatin that includes regulatory sequences and transposable elements.383–385 As EGA approaches, accessible chromatin becomes more structured, leading to the detection of an increasing number of consistently open chromatin regions at specific locations but an overall decrease in global chromatin accessibility.384–386 There is a subsequent progressive increase in the number of specific regions of accessible chromatin during human preimplantation embryo development from the 2-­cell to blastocyst stages.385 Changes in chromatin accessibility during preimplantation development are accompanied by progressive maturation of chromatin structure by the formation of 3-­dimensional “topologically associating domains” (TADs). TADs consist of loops of chromatin that maintain gene promoters and distal regulatory elements in close proximity and are important for regulating gene expression.387–389 TADs are established through DNA interactions with the cohesion complex and CTCF (CCCTC-­binding factor), a transcriptional repressor. In human embryos, TADs begin to be established at the time of EGA at the 8-­cell stage and are fully developed at the blastocyst stage.389 Knockdown of CTCF in human zygotes prevents TAD establishment, emphasizing the importance of this protein in regulating embryonic gene expression. Maternal proteins generated from stored mRNAs contribute to the regulation of embryonic gene expression through impacts on chromatin structure. Most of the information regarding maternally inherited factors that impact gene expression has been gleaned from studies in mouse models. For example, SMARCA4 (previously known as BRG1) is the catalytic subunit

of a chromatin remodeling complex that alters nucleosome conformation and positioning, increasing chromatin accessibility. In the mouse, SMARCA4 is maternally inherited and when eggs lacking SMARCA4 are fertilized they fail to undergo EGA.390,391 SIN3A, a scaffolding protein that binds histone deacetylases and functions as a transcriptional corepressor, is also maternally inherited in the mouse.392 Embryos lacking SIN3A have alterations in histone acetylation and gene transcription and cannot develop beyond the 2-­cell stage. Although these examples are from mice, it is highly likely that human embryos also inherit maternal proteins that promote chromatin remodeling based on the changes in chromatin structure that occur prior to EGA at the 8-­cell stage. Chromatin remodeling is permissive for EGA, but it is not sufficient because there is a need for transcription factors to direct the transcriptional machinery to the correct locations. One important maternally inherited transcription factor is nuclear transcription factor Y (NFY). NFY is a pioneer factor responsible for maintaining the nucleosome depletion of promoter regions so that the transcriptional machinery can reach specific transcription start sites.393 Mouse embryos lacking NFY do not open chromatin at the appropriate promoter regions at the 2-­cell stage and fail to develop beyond the morula stage.394 ß-­catenin is a maternally inherited transcription factor that functions in the mouse zygote and 2-­cell stage embryo to promote the transcription of genes required for ribosomal biogenesis.142 Without ß-­catenin function, the early embryo fails to synthesize new proteins necessary for development beyond the 2-­cell stage. Embryo-­derived transcription factors are also essential for activating gene expression required for EGA. The earliest expressed embryonic transcription factor identified to date is double homeobox 4 (DUX4), which was identified by analyzing the promoters of genes encoding transcripts present in cleavage stage human embryos.395,396 A DNA motif predicted to bind DUX4 was highly enriched in these genes and further experiments demonstrated that DUX4 activates expression of cleavage stage genes, including other transcription factors and chromatin remodeling proteins. Interestingly, DUX4 also activates expression of repetitive elements including the endogenous retrovirus MERVL; expression of repetitive elements is thought to promote chromatin accessibility in the cleavage stage embryo. DUX, the mouse homolog of human DUX4, has a similar role in EGA. Expression of mouse Dux and human Dux4 is likely initiated by multiple inputs. Both of these genes are nested within large repetitive DNA sequences that are normally suppressed in regions of heterochromatin.397 Global chromatin relaxation as a consequence of paternal protamine/histone exchange and activity of chromatin remodelers in the zygote and early cleavage stage embryo may lead to de-­repression of Dux and Dux4. There is evidence in the mouse that two maternally inherited DNA-­ binding proteins, developmental pluripotency-­ associated gene 2 (DPPA2) and DPPA4, activate Dux expression.398,399 DPPA2 and DPPA4 also induce expression of LINE-­1 retrotransposons in early embryos, which may contribute to chromatin relaxation. DUX itself promotes expression of Dppa2, so rather than a simple linear series of transcription factors activating downstream factors, EGA appears to require multiple inputs and feedback loops for its success.

COMPACTION AND BLASTOCYST DEVELOPMENT • C  ompaction depends on the establishment of E-­cadherin-­based adherens junctions between outer cells of the morula. • Hippo and Notch signaling drive formation of the first distinct cell lineages at the morula stage: the outer cells become trophectoderm and the inner cells become inner cell mass cells. • Tight junctions are formed in trophectoderm cells and are essential

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development Compacted

SEM

DIC

Noncompacted

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domains to establish adherens junctions between the blastomeres. This homotypic E-­ cadherin binding is extracellular calcium-­ dependent, explaining why incubation of embryos in a calcium-­ free medium facilitates blastomere biopsy. Additional cell–cell connections are achieved by the extension of filopodia from one blastomere over the surface of adjacent blastomeres; these filopodia depend on the presence of E-­cadherin.404 Several important changes to the apical blastomere surface occur during compaction. The actin-­associated protein, ezrin, becomes localized to the apical surface and is no longer observed at cell–cell junctions.405 Ezrin has a critical role in the formation of apical microvilli.406 In addition, the actomyosin cytoskeleton becomes highly enriched in the central region of the apical surface. Periodic contractions of this apical actomyosin cortex appear to provide the force that drives both compaction and cell internalization as the blastomeres continue to divide.407 A consequence of cell internalization is the formation of morphologically distinct cell lineages, outer cells, and inner cells, for the first time in the morula stage embryo.

Morula to Blastocyst Transition

Fig. 9.21 Morphological changes in mouse preimplantation embryos undergoing compaction.  Membranes of individual blastomeres are clearly delineated under DIC optics in a noncompacted 8-­cell mouse embryo (top left panel). Scanning electron microscopy reveals a uniform distribution of microvilli across all cell surfaces and blastomeres are relatively spherical (bottom left panel). Individual cell membranes are no longer discernible by DIC in the compacted embryo (top right panel). The blastomeres have flattened and the microvilli are localized to apical zones surrounded by a smooth membrane at cell–cell junctions (bottom right panel). Scale bars: 10 μm x, y and 15 μm x, y, z. (From White MD, Bissiere S, Alvarez YD, Plachta N. Mouse embryo compaction. Curr Top Dev Biol. 2016;120:235–258. https://doi.org/10.1016/bs.ctdb.2016.04.005.)

• B  lastocoel formation is a consequence of the active transport of ions across the outer cells followed by passive entry of water due to the resulting osmotic gradient. • Development of more than one inner cell mass in the blastocyst can cause monozygotic twinning. • The blastocyst expands in size and hatches from the zona pellucida prior to implantation.

Compaction At the end of the cleavage stage, the individual blastomeres become tightly associated with each other in a process known as compaction. Compaction changes the appearance of the embryo from a collection of relatively spherical, nonpolar blastomeres to a tightly packed group of polarized cells that lack visible cell boundaries or intercellular spaces (Fig. 9.21). In both mouse and human embryos, compaction usually occurs during the 8-­to 16-­ cell stages.400,401 After compaction, the embryo is termed a morula. Compaction relies on cell–cell interactions mediated by the cell adhesion molecule, E-­cadherin. In early cleavage stage embryos, E-­cadherin is expressed uniformly around all regions of blastomere plasma membranes. It is connected to the subcortical actin cytoskeleton through interactions of its intracellular domain with α-­ catenin and ß-­catenin, which functions in the actin cytoskeleton in addition to its role as a transcription factor. At the 4-­to 8-­cell stage, E-­cadherin, ß-­catenin, and α-­catenin become restricted to cell–cell junctions.402 Compaction is likely initiated by the action of protein kinase C-­alpha, which translocates to membranes of cell–cell junctions and phosphorylates ß-catenin.403 E-cadherin molecules on adjacent cells bind to each other through their extracellular

The morula stage (∼16-­to 32-­cells) is characterized by an increase in the numbers of inner nonpolar blastomeres and outer blastomeres that undergo a gradual conversion to a true epithelium that has apical-­basal polarity and tight junctions. Tight junctions are located at epithelial cell–cell junctions at the apical end of lateral membranes, where they form a barrier that prevents entry of solutes and macromolecules into paracellular spaces. The embryo forms tight junctions by first incorporating tight junction protein 1 (TJP1, formerly zona occludens 1) and cingulin in locations of adherens junctions.408 RAB13, a GTPase that regulates vesicular transport of proteins to the membrane, is also found in the same locations, suggesting that RAB13 regulates tight junction complex assembly by controlling the delivery of constituent proteins. Finally, occludin and claudins, transmembrane proteins that support cell–cell adhesion through homotypic interactions, become incorporated into a complex with TJP1, cingulin, and other actin cytoskeleton-­linked proteins, forming a functional tight junction.409,410 Mature tight junctions cause close apposition of the adjacent cell membranes and formation of a functional diffusion barrier that supports the transition to the blastocyst stage (Fig. 9.22). Cavitation, or formation of the fluid-­filled blastocoel (blastocyst cavity), marks the beginning of the blastocyst stage (Fig. 9.23). Cavitation only occurs following the formation of functional tight junctions and is caused by active transepithelial transport of sodium and chloride ions, resulting in an ionic gradient across the trophectoderm.411 In response to the ionic gradient, water is passively transported across the trophectoderm into the inner cell mass through aquaporin water channels located on both the apical and basolateral membranes.412 Cavitation is regulated by at least two different kinases, PKA and MAPK14 (p38 mitogen-­ associated protein kinase).413,414 Collections of fluid inside the early blastocyst eventually coalesce to form a single cavity, leaving a group of nonpolar inner cell mass (ICM) cells eccentrically located adjacent to the outer polar cells, the trophectoderm. The ICM will eventually form the embryo proper and the trophectoderm will form the fetal cells of the placenta.

Monozygotic Twinning Most twins are a result of dizygotic twinning, which occurs when two different eggs are fertilized following polyovulation (see Chapter 8). Monozygotic twins, a phenomenon regularly observed only in humans and one other vertebrate, the armadillo, develop from a single fertilized egg and represent about ∼30% of liveborn spontaneous human twins.415 There are several theories regarding how and when during development monozygotic twins form. A long-­ held idea

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Undifferentiated blastomeres Trophectoderm lineage ICM lineage

Pre-compact 8-cell

Compact 8-cell

16-cell morula

Early blastocyst ~32-cell

ZO-1α– RAB13 cingulin ZO-1α+ occludin E-cadherin/ catenin

Permeable

Impermeable

Fig. 9.22 Schematic diagram of preimplantation stages and proposed pattern of membrane assembly of junctional proteins during trophectoderm differentiation.  Below each stage is shown the composition of apicolateral contact sites between outer cells. Following the assembly of occludin at the 32-­cell stage, separation of apical tight junction and subjacent zonula adherens/E-­cadherin domains is apparent, coinciding with blastocyst cavitation and the establishment of a permeability seal. (From Sheth B, Fontaine JJ, Ponza E, et al. Differentiation of the epithelial apical junctional complex during mouse preimplantation development: a role for rab13 in the early maturation of the tight junction. Mech Dev. 2000;97(1–2):93–104. https://doi.org/10.1016/S0925-4773(00)00416-0.)

A

B

C

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E

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H

Fig. 9.23 Development of human embryos between 90 and 120 hours after insemination.  (A) A morula with a characteristic loss of cell borders define a cleavage stage embryo. The zona pellucida is still thick (arrowhead). (B) A morula transitioning to a very early blastocyst at 92 hours; note that cavitation results in the accumulation of small pockets of blastocoel fluid (arrow), and the first overt signs of cellular differentiation with epithelial cells forming at the periphery that will develop into the trophectoderm cells. (C) An early blastocyst at 100 hours; note the increasing size of fluid pockets (arrow). (D) Further differentiation of the trophectoderm cells, and the appearance of a small cluster of spherical cells that will form the inner cell mass. (E) A fully expanded blastocyst at 116 hours; note the thinned zona pellucida and the initial herniation of trophectoderm cells. (F) A blastocyst undergoing contraction, which is followed by subsequent re-­ expansion. The trophectoderm is now a well-organized epithelial layer ( row). (G) A hatched blastocyst with a compact inner cell mass ( trophectoderm (arrowhead). (H) Zona pellucida corresponding to hatched blastocyst in (G) containing discarded cytoplasmic fragments (arrow).

was that a zygote could split into more than one embryo within the zona pellucida during development to the morula stage, eventually resulting in monozygotic, dichorionic/diamniotic twins, each supported by a distinct placenta. However, the absence of any reports of embryo splitting during in vitro culture of human embryos to date, despite the millions of embryos observed worldwide, indicates that this mechanism is highly unlikely.416,417 In contrast, there is good evidence from live imaging of human preimplantation embryo development during assisted reproduction cycles that two distinct ICMs sometimes form (Fig. 9.24).418 If both ICMs develop into embryos without splitting the surrounding trophoblast, then monozygotic, monochorionic/diamniotic twins will result. Interestingly, ICM splitting was documented in one case to result in a monochorionic/triamniotic triplet pregnancy following the transfer of a single embryo.419 Should the trophoblast cells also split into distinct layers, one surrounding each ICM, then monozygotic, dichorionic/diamniotic twins can result. This type of monozygotic twinning is less common, representing about 30% of liveborn monozygotic twin cases.420 A convincing demonstration of this type of twinning was reported in several cases following single frozen embryo transfer in controlled, downregulated cycles in which the possibility of a concomitant spontaneous pregnancy was remote.421 The rare cases of monozygotic, monochorionic/monoamniotic twinning (1%-­2% of liveborn monozygotic twins) result when embryo splitting occurs following implantation and formation of the amnion.415 The mechanism(s) underlying the splitting of the ICM or trophoblast cells is not known, but there is human genetic support for the idea that a diminished capacity for cell adhesion could contribute. Although it is a rare phenomenon, there are reports of familial monozygotic twinning. A recent study of a four-­generation family with seven pairs of monozygotic twins used whole genome sequencing to identify potential contributing genetic factors.422 Analysis of both single nucleotide variants and copy number variants suggested that the identified genes were most significantly enriched in two pathways relevant to cell adhesion: epithelial adherens junction and tight junction signaling. These findings support the idea that impaired cell adhesion could be responsible for monozygotic twinning.

Formation of Distinct Cell Lineages Cell lineage specification is a gradual process that begins after the 8-cell stage. In mice, Hippo signaling, a mechanism of controlling cell proliferation that depends on cell polarity and cell–cell contacts, regulates the initial specification of trophectoderm and ICM.423 Mouse early morulas express several important lineage-determining

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development

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Fig. 9.24 Two apparent inner cell masses in single blastocyst observed during a retrospective review of live embryo imaging.  Single blastocyst transfer resulted in monozygotic, monochorionic/triamnionic triplet pregnancy. Panels are the same embryo in different focal planes. Boxes indicate inner cell masses. (Modified from Sutherland K, Leitch J, Lyall H, Woodward BJ. Time-­lapse imaging of inner cell mass splitting with monochorionic triamniotic triplets after elective single embryo transfer: a case report. Reprod Biomed Online. 2019;38(4):491–496. https://doi.org/10.1016/j.rbmo.2018.12.017.)

transcription factors in most cells at similar levels. These factors include POU5F1 (POU domain class 5 transcription factor 1, formerly known as OCT4), CDX2 (caudal-­type homeobox transcription factor 2), NANOG (homeobox transcription factor NANOG), TEAD4 (TEA domain factor 4), and the transcriptional coactivator YAP (YES-­associated protein). The nonpolar inner cells have extensive cell–cell contacts that cause activation of Hippo signaling at adherens junctions, leading to phosphorylation and degradation of YAP. In contrast, the polarized outer cells have fewer cell–cell contacts and do not activate Hippo signaling, allowing YAP to translocate to the nucleus. Although TEAD4 is expressed in all cells of the embryo, it is not active in the absence of YAP. When YAP translocates to the nucleus of outer cells it functions together with TEAD4 to activate the transcription of Cdx2, a driver of trophectoderm differentiation.424,425 Rising CDX2 levels suppress transcription of Pou5f1, eventually restricting POU5F1 to the ICM.426 Reciprocally, increased levels of POU5F1 in the ICM suppress transcription of Cdx2, preventing differentiation of the ICM into trophoblast cells. Notch signaling, another cell contact-­driven signaling pathway, also contributes to lineage specification of trophoblast cells by promoting the expression of Cdx2 in outer cells.427 In this way, positional information obtained from cell–cell contact-­ mediated signaling allows the embryo to self-­organize into two distinct cell lineages, the trophectoderm and the ICM. Lineage specification in human embryos appears to occur in the same way as in mice except that the timing is somewhat delayed. Human 8-­cell embryos have essentially the same transcriptional profile in each blastomere and they contain transcripts encoding many lineage-­specific transcription factors.428 However, exactly when the proteins are generated is only well documented in a few cases. For example, CDX2 is expressed in most cells in early blastocysts but becomes restricted to the trophectoderm cells in late blastocysts (>75 cells).429 Similarly, POU5F1 is expressed in both trophectoderm and ICM cells until the late blastocyst stage when it becomes restricted to the ICM. Restriction of these transcription factors to specific cell lineages may occur later than in the mouse because of the relatively longer time of preimplantation embryo development. Slower differentiation of the ICM could contribute to the occurrence of split ICMs and monozygotic twinning in human embryos. Before implantation, the ICM itself differentiates into two distinct cell lineages: the primitive endoderm, which contributes

cells to the extraembryonic yolk sac and visceral endoderm, and the epiblast, which will become the embryo proper and contributes cells to the extra-­ embryonic mesoderm (Fig. 9.25). The primitive endoderm lies adjacent to the blastocoel and the epiblast is located in between the primitive endoderm and trophectoderm cells. All ICM cells initially express POU5F1, but they also express either of two additional transcription factors, NANOG or GATA6, in a “salt and pepper” pattern.430,431 These cells sort themselves into two distinct cell layers separated by a basal lamina under the influence of intraembryonic mitogen-­associated protein kinase and fibroblast growth factor 4 signaling.431,432 At this point, the epiblast cells express NANOG and the primitive endoderm cells express GATA6.431 Additional complexity in the process of ICM differentiation is conferred by the activation of Hippo/YAP signaling in the ICM of later-­stage blastocysts.433 YAP and TEAD together induce high-­ level expression of factors important for pluripotency, which is critical for the ability of the ICM to generate most embryonic cell types.

Blastocyst Expansion and Hatching As cell proliferation and cell lineage specification continues, the blastocoel enlarges, leading eventually to an increase in the overall diameter of the blastocyst. This process is known as “blastocyst expansion” and is accompanied by zona pellucida thinning and increased susceptibility to protease digestion. Interestingly, blastocyst expansion does not occur progressively; instead, embryos have periodic oscillations in their expansion activity.434 During the blastocyst expansion phase, trophectoderm cells form desmosomes, which are highly adhesive cell–cell junctions that stabilize epithelia that are subject to mechanical stresses.435 In theory, inadequate desmosome formation could contribute to the disruption of trophectoderm continuity and promote formation of monozygotic, dichorionic/diamniotic twins if ICM splitting also occurs. The expanded blastocyst eventually “hatches” from the zona pellucida in preparation for implantation. Hatching is a gradual process mediated by blastocyst-­derived proteases and prostaglandins and is promoted by the presence of growth factors secreted The hatched blastocyst is poised to interact directly with the receptive endometrium and begin the process of implantation.

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TG

ICM

ZGA/EGA

ExEc ExEn Epi

Epi PE TE

Mouse Pronuclei

Post-implantation

Preimplantation

Syn

VCT Embryo

ICM

EGA

Yolk sac

Human Preimplantation Post-implantation Embryonic 0 day

1

2

3

4

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6

Fig. 9.25 Cell fate decisions and their timing in mouse versus human early embryo development.  Prior to implantation, both human and mouse embryos similarly undergo cell divisions culminating in the development of a blastocyst comprising a discernible inner cell mass (ICM) and trophectoderm (TE). Mouse zygotic/embryonic genome activation (ZGA/EGA) begins at the 2-­cell stage, whereas human EGA begins at the ∼4-­to 8-­cell stage on day 3, although minor human EGA may occur as early as the 2-­cell stage. The timing of compaction and blastocyst formation also differs significantly, with human embryos showing delayed development compared with mouse embryos; the mouse blastocyst forms between days 3 and 4, whereas human blastocysts form between days 5 and 6. Both human and mouse pre-­implantation blastocysts comprise an outer layer of TE cells (blue), which form the trophoblast lineage of the placenta, and an ICM that segregates into epiblast (Epi, green) and primitive endoderm (PE, red) layers. Epiblast cells eventually give rise to all the tissues of the future fetus, whereas the PE gives rise to extra-­embryonic endoderm (ExEn) cells that will form the yolk sac. In the mouse, the TE gives rise to a proliferative stem cell pool of extra-­embryonic ectoderm (ExEc) cells that bud off differentiated polyploid trophoblast giant (TG) cells. By contrast, human TE gives rise to villous cytotrophoblast (VCT) cells, a multinucleated syncytium (Syn), and extravillous trophoblast cells (not shown). The dashed arrow indicates minor gene activation. (Modified from Niakan KK, Han J, Pedersen RA, Simon C, Pera RA. Human pre-­implantation embryo development. Development. 2012;139(5):829–841. https://doi.org/10.1242/dev.060426.)

TOP REFERENCES

Bhakta HH, Refai FH, Avella MA. The molecular mechanisms mediating mammalian fertilization. Development. 2019;146:dev176966. Available at: http://dev.biologists.org/lookup/doi/10.1242/dev.176966. Clift D, Schuh M. A three-step MTOC fragmentation mechanism facilitates bipolar spindle assembly in mouse oocytes. Nat Commun. 2015;6:7217. Available at: http://www.ncbi.nlm.nih.gov/ pubmed/26147444. Eckersley-Maslin MA, Alda-Catalinas C, Reik W. Dynamics of the epigenetic landscape during the maternal-to-zygotic transition. Nat Rev Mol Cell Biol. 2018;19:436–450. Available at: https://doi.org/10.1038/ s41580-018- 0008-z. Giojalas LC, Guidobaldi HA. Getting to and away from the egg, an interplay between several sperm transport mechanisms and a complex oviduct physiology. Mol Cell Endocrinol. 2020;518:110954. Gruhn JR, Zielinska AP, Shukla V, et al. Chromosome errors in human eggs shape natural fertility over reproductive life span. Science. 2019;365:1466–1469. Available at: https://www.ncbi.nlm.nih.gov/ pubmed/31604276. Halstead MM, Ma X, Zhou C, Schultz RM, Ross PJ. Chromatin remodeling in bovine embryos indicates species-specific regulation of genome activation. Nat Commun. 2020:11. Available at: https://doi. org/10.1038/s41467-020-18508-3. Ishiguro KI, Matsuura K, Tani N, et al. MEIOSIN directs the switch from mitosis to meiosis in mammalian germ cells. Dev Cell. 2020;52:429–445.e10. Available at: https://www.ncbi.nlm.nih.gov/ pubmed/32032549.

Mehlmann LM, Saeki Y, Tanaka S, et al. The Gs-linked receptor GPR3 maintains meiotic arrest in mammalian oocytes. Science. 2004;306:1947–1950. Available at: http://www.ncbi.nlm.nih.gov/ entrez/query.fcgi?cmd=Retrieve&db=PubMed&dopt=Citation&l ist_uids=15591206. Niakan KK, Han J, Pedersen RA, Simon C, Pera RAR. Human preimplantation embryo development. Development. 2012;139:829–841. Stein P, Savy V, Williams AM, Williams CJ. Modulators of calcium signalling at fertilization. Open Biol. 2020;10. Tokuhiro K, Dean J. Glycan-independent gamete recognition triggers egg zinc sparks and ZP2 cleavage to prevent polyspermy. Dev Cell. 2018;46:627–640. e5. Available at: https://doi.org/10.1016/j. devcel.2018.07.020. Trebichalská Z, Kyjovská D, Kloudová S, Otevřel P, Hampl A, Holubcová Z. Cytoplasmic maturation in human oocytes: an ultrastructural study †. Biol Reprod. 2020;104:106–116. Vastenhouw NL, Cao WX, Lipshitz HD. The maternal-to-zygotic transition revisited. Development. 2019:146. Wang H, McGoldrick LL, Chung J-J. Sperm ion channels and transporters in male fertility and infertility. Nat Rev Urol. 2021;18:46–66. Available at: https://doi.org/10.1038/s41585-020-00390-9. White MD, Plachta N. Specification of the first mammalian cell lineages in vivo and in vitro. Cold Spring Harb Perspect Biol. 2020;12:1–24.

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REFERENCES

1. Mehlmann LM. Stops and starts in mammalian oocytes: recent advances in understanding the regulation of meiotic arrest and oocyte maturation. J Reprod Fertil. 2005;130(6):791–799. 2. Endo T, Mikedis MM, Nicholls PK, Page DC, de Rooij DG. Retinoic acid and germ cell development in the ovary and testis. Biomolecules. 2019;9(12):775. 3. Childs AJ, Cowan G, Kinnell HL, Anderson RA, Saunders PTK. Retinoic acid signalling and the control of meiotic entry in the human fetal gonad. PLoS One. 2011;6(6):e20249. 4. Jørgensen A, Nielsen JE, Blomberg Jensen M, Græm N, Rajpert-­De Meyts E. Analysis of meiosis regulators in human gonads: a sexually dimorphic spatio-­temporal expression pattern suggests involvement of DMRT1 in meiotic entry. Mol Hum Reprod. 2012;18(11):523–534. 5. Le Bouffant R, Guerquin MJ, Duquenne C, et al. Meiosis initiation in the human ovary requires intrinsic retinoic acid synthesis. Hum Reprod. 2010;25(10):2579–2590. 6. Baltus AE, Menke DB, Hu YC, et al. In germ cells of mouse embryonic ovaries, the decision to enter meiosis precedes premeiotic DNA replication. Nat Genet. 2006;38(12):1430–1434. 7. Koubova J, Menke DB, Zhou Q, Capel B, Griswold MD, Page DC. Retinoic acid regulates sex-­specific timing of meiotic initiation in mice. Proc Natl Acad Sci U S A. 2006;103(8):2474–2479. 8. Feng CW, Burnet G, Spiller CM, et al. Identification of regulatory elements required for stra8 expression in fetal ovarian germ cells of the mouse. Development. 2021;148(5):dev194977. 9. Mark M, Jacobs H, Oulad-­Abdelghani M, et al. STRA8-­deficient spermatocytes initiate, but fail to complete, meiosis and undergo premature chromosome condensation. J Cell Sci. 2008;121(Pt 19):3233–3242. 10. Gill ME, Hu YC, Lin Y, Page DC. Licensing of gametogenesis, dependent on RNA binding protein DAZL, as a gateway to sexual differentiation of fetal germ cells. Proc Natl Acad Sci U S A. 2011;108(18):7443–7448. 11. Lin Y, Page DC. Dazl deficiency leads to embryonic arrest of germ cell development in XY C57BL/6 mice. Dev Biol. 2005;288(2):309–316. 12. Ishiguro KI, Matsuura K, Tani N, et al. MEIOSIN directs the switch from mitosis to meiosis in mammalian germ cells. Dev Cell. 2020;52(4):429–445.e10. 13. Li H, MacLean G, Cameron D, Clagett-­Dame M, Petkovich M. Cyp26b1 expression in murine Sertoli cells is required to maintain male germ cells in an undifferentiated state during embryogenesis. PLoS One. 2009;4(10):e7501. 14. MacLean G, Li H, Metzger D, Chambon P, Petkovich M. Apoptotic extinction of germ cells in testes of Cyp26b1 knockout mice. Endocrinology. 2007;148(10):4560–4567. 15. Hogarth CA, Evans E, Onken J, et al. CYP26 enzymes are necessary within the postnatal seminiferous epithelium for normal murine spermatogenesis. Biol Reprod. 2015;93(1):19. 16. Hogarth CA, Evanoff R, Snyder E, et al. Suppression of Stra8 expression in the mouse gonad by WIN 18,446. Biol Reprod. 2011;84(5):957–965. 17. Wolbach SB, Howe PR. Tissue changes following deprivation of fat-­soluble a vitamin. J Exp Med. 1925;42(6):753–777. 18. Romanienko PJ, Camerini-­ Otero RD. The mouse Spo11 gene is required for meiotic chromosome synapsis. Mol Cell. 2000;6(5):975–987. 19. Grey C, Barthès P, Chauveau-­Le Friec G, Langa F, Baudat F, de Massy B. Mouse PRDM9 DNA-­ binding specificity determines sites of histone H3 lysine 4 trimethylation for initiation of meiotic recombination. PLoS Biol. 2011;9(10):e1001176. 20. Brick K, Smagulova F, Khil P, Camerini-­Otero RD, Petukhova GV. Genetic recombination is directed away from functional genomic elements in mice. Nature. 2012;485(7400):642–645. 21. Baudat F, Buard J, Grey C, et al. PRDM9 is a major determinant of meiotic recombination hotspots in humans and mice. Science. 2010;327(5967):836–840. 22. Billings T, Parvanov ED, Baker CL, Walker M, Paigen K, Petkov PM. DNA binding specificities of the long zinc-­finger recombination protein PRDM9. Genome Biol. 2013;14(4):R35. 23. Walker M, Billings T, Baker CL, et al. Affinity-­seq detects genome-­ wide PRDM9 binding sites and reveals the impact of prior chromatin modifications on mammalian recombination hotspot usage. Epigenet Chromatin. 2015;8(1):31.

24. Powers NR, Parvanov ED, Baker CL, Walker M, Petkov PM, Paigen K. The meiotic recombination activator PRDM9 trimethylates both H3K36 and H3K4 at recombination hotspots in vivo. PLoS Genet. 2016;12(6):e1006146. 25. Hayashi K, Yoshida K, Matsui Y. A histone H3 methyltransferase controls epigenetic events required for meiotic prophase. Nature. 2005;438(7066):374–378. 26. Narasimhan VM, Hunt KA, Mason D, et al. Health and population effects of rare gene knockouts in adult humans with related parents. Science. 2016;352(6284):474–477. 27. Irie S, Tsujimura A, Miyagawa Y, et al. Single-­nucleotide polymorphisms of the PRDM9 (MEISETZ) gene in patients with nonobstructive azoospermia. J Androl. 2009;30(4):426–431. 28. Powers NR, Dumont BL, Emori C, et al. Sexual dimorphism in the meiotic requirement for PRDM9: a mammalian evolutionary safeguard. Sci Adv. 2020;6(43):eabb6606. 29. Biswas L, Tyc K, El Yakoubi W, Morgan K, Xing J, Schindler K. Meiosis interrupted: the genetics of female infertility via meiotic failure. J Reprod Fertil. 2021;161(2):R13–R35. 30. Geisinger A, Benavente R. Mutations in genes coding for synaptonemal complex proteins and their impact on human fertility. Cytogenet Genome Res. 2016;150(2):77–85. 31. Bolor H, Mori T, Nishiyama S, et al. Mutations of the SYCP3 gene in women with recurrent pregnancy loss. Am J Hum Genet. 2009;84(1):14–20. 32. Sazegari A, Kalantar SM, Pashaiefar H, et al. The T657C polymorphism on the SYCP3 gene is associated with recurrent pregnancy loss. J Assist Reprod Genet. 2014;31(10):1377–1381. 33. McGuire MM, Bowden W, Engel NJ, Ahn HW, Kovanci E, Rajkovic A. Genomic analysis using high-­resolution single-­nucleotide polymorphism arrays reveals novel microdeletions associated with premature ovarian failure. Fertil Steril. 2011;95(5):1595–1600. 34. de Vries L, Behar DM, Smirin-­Yosef P, Lagovsky I, Tzur S, Basel-­ Vanagaite L. Exome sequencing reveals SYCE1 mutation associated with autosomal recessive primary ovarian insufficiency. J Clin Endocrinol Metab. 2014;99(10):E2129–E2132. 35. Hernández-­López D, Geisinger A, Trovero MF, et al. Familial primary ovarian insufficiency associated with an SYCE1 point mutation: defective meiosis elucidated in humanized mice. Mol Hum Reprod. 2020;26(7):485–497. 36. Li XC, Schimenti JC. Mouse pachytene checkpoint 2 (trip13) is required for completing meiotic recombination but not synapsis. PLoS Genet. 2007;3(8):e130. 37. Zhang Z, Li B, Fu J, et al. Bi-­allelic missense pathogenic variants in TRIP13 cause female infertility characterized by oocyte maturation arrest. Am J Hum Genet. 2020;107(1):15–23. 38. Kauppi L, Barchi M, Baudat F, Romanienko PJ, Keeney S, Jasin M. Distinct properties of the XY pseudoautosomal region crucial for male meiosis. Science. 2011;331(6019):916–920. 39. Royo H, Polikiewicz G, Mahadevaiah SK, et al. Evidence that meiotic sex chromosome inactivation is essential for male fertility. Curr Biol. 2010;20(23):2117–2123. 40. Cloutier JM, Mahadevaiah SK, ElInati E, Tóth A, Turner J. Mammalian meiotic silencing exhibits sexually dimorphic features. Chromosoma. 2016;125(2):215–226. 41. Lutzmann M, Grey C, Traver S, et al. MCM8-­and MCM9-­deficient mice reveal gametogenesis defects and genome instability due to impaired homologous recombination. Mol Cell. 2012;47(4):523–534. 42. Wood-­Trageser MA, Gurbuz F, Yatsenko SA, et al. MCM9 mutations are associated with ovarian failure, short stature, and chromosomal instability. Am J Hum Genet. 2014;95(6):754–762. 43. Bolcun-­Filas E, Rinaldi VD, White ME, Schimenti JC. Reversal of female infertility by Chk2 ablation reveals the oocyte DNA damage checkpoint pathway. Science. 2014;343(6170):533–536. 44. Rinaldi VD, Bolcun-­Filas E, Kogo H, Kurahashi H, Schimenti JC. The DNA damage checkpoint eliminates mouse oocytes with chromosome synapsis failure. Mol Cell. 2017;67(6):1026–1036.e2. 45. Rinaldi VD, Bloom JC, Schimenti JC. Oocyte elimination through DNA damage signaling from CHK1/CHK2 to p53 and p63. Genetics. 2020;215(2):373–378. 46. Giacomo D, Barchi M, Baudat M, Edelmann F, Keeney W, Jasin S. Distinct DNA-­damage-­dependent and -­independent responses drive the loss of oocytes in recombination-­defective mouse mutants. Proc Natl Acad Sci U S A. 2005;102:737–742.

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216.e2 PART I  The Fundamentals of Reproduction 47. Marcet-­Ortega M, Pacheco S, Martínez-­Marchal A, et al. p53 and TAp63 participate in the recombination-­ dependent pachytene arrest in mouse spermatocytes. PLoS Genet. 2017;13(6):e1006845. 48. Jan SZ, Jongejan A, Korver CM, et al. Distinct prophase arrest mechanisms in human male meiosis. Development. 2018; 145(16):dev160614. 49. de Boer E, Stam P, Dietrich AJJ, Pastink A, Heyting C. Two levels of interference in mouse meiotic recombination. Proc Natl Acad Sci U S A. 2006;103(25):9607–9612. 50. Kolas NK, Svetlanov A, Lenzi ML, et al. Localization of MMR proteins on meiotic chromosomes in mice indicates distinct functions during prophase I. J Cell Biol. 2005;171(3):447–458. 51. Eaker S, Cobb J, Pyle A, Handel MA. Meiotic prophase abnormalities and metaphase cell death in MLH1-­deficient mouse spermatocytes: insights into regulation of spermatogenic progress. Dev Biol. 2002;249(1):85–95. 52. Edelmann W, Cohen PE, Kane M, et al. Meiotic pachytene arrest in MLH1-­deficient mice. Cell. 1996;85(7):1125–1134. 53. Lipkin SM, Moens PB, Wang V, et al. Meiotic arrest and aneuploidy in MLH3-­deficient mice. Nat Genet. 2002;31(4):385–390. 54. Markandona O, Dafopoulos K, Anifandis G, et al. Single-­nucleotide polymorphism rs 175080 in the MLH3 gene and its relation to male infertility. J Assist Reprod Genet. 2015;32(12):1795–1799. 55. Pashaiefar H, Sheikhha MH, Kalantar SM, Jahaninejad T, Zaimy MA, Ghasemi N. Analysis of MLH3 C2531T polymorphism in Iranian women with unexplained infertility. Iran J Reprod Med. 2013;11(1):19–24. 56. Singh P, Fragoza R, Blengini CS, et al. MLH1/3 Variants Causing Aneuploidy, Pregnancy Loss, and Premature Reproductive Aging. bioRxiv; 2021. Published online. 57. Wesley ER, Hawley RS, Billmyre KK. Genetic background impacts the timing of synaptonemal complex breakdown in drosophila melanogaster. Chromosoma. 2020;129(3–4):243–254. 58. Wellard SR, Schindler K, Jordan PW. Aurora B and C kinases regulate chromosome desynapsis and segregation during mouse and human spermatogenesis. J Cell Sci. 2020;133(23):jcs248831. 59. Guan Y, Leu NA, Ma J, et al. SKP1 drives the prophase I to metaphase I transition during male meiosis. Sci Adv. 2020;6(13): eaaz2129. 60. De La Fuente R. Chromatin modifications in the Germinal Vesicle (GV) of mammalian oocytes. Dev Biol. 2006;292(1):1–12. 61. De La Fuente R, Viveiros MM, Burns KH, Adashi EY, Matzuk MM, Eppig JJ. Major chromatin remodeling in the Germinal Vesicle (GV) of mammalian oocytes is dispensable for global transcriptional silencing but required for centromeric heterochromatin function. Dev Biol. 2004;275(2):447–458. 62. Johnson LD, Albertini DF, McGinnis LK, Biggers JD. Chromatin organization, meiotic status and meiotic competence acquisition in mouse oocytes from cultured ovarian follicles. J Reprod Fertil. 1995;104(2):277–284. 63. Zuccotti M, Boiani M, Garagna S, Redi CA. Analysis of aneuploidy rate in antral and ovulated mouse oocytes during female aging. Mol Reprod Dev. 1998;50(3):305–312. 64. Bonnet-­Garnier A, Feuerstein P, Chebrout M, et al. Genome organization and epigenetic marks in mouse germinal vesicle oocytes. Int J Dev Biol. 2012;56(10–12):877–887. 65. Bornslaeger EA, Mattei P, Schultz RM. Involvement of cAMP-­ dependent protein kinase and protein phosphorylation in regulation of mouse oocyte maturation. Dev Biol. 1986;114(2):453–462. 66. Cho WK, Stern S, Biggers JD. Inhibitory effect of dibutyryl cAMP on mouse oocyte maturation in vitro. J Exp Zool. 1974;187(3): 383–386. 67. Magnusson C, Hillensjö T. Inhibition of maturation and metabolism in rat oocytes by cyclic amp. J Exp Zool. 1977;201(1):139–147. 68. Vaccari S, Weeks 2nd JL, Hsieh M, Menniti FS, Conti M. Cyclic GMP signaling is involved in the luteinizing hormone-­ dependent meiotic maturation of mouse oocytes. Biol Reprod. 2009;81(3):595–604. 69. Hinckley M, Vaccari S, Horner K, Chen R, Conti M. The G-­protein-­ coupled receptors GPR3 and GPR12 are involved in cAMP signaling and maintenance of meiotic arrest in rodent oocytes. Dev Biol. 2005;287(2):249–261. 70. Horner K, Livera G, Hinckley M, Trinh K, Storm D, Conti M. Rodent oocytes express an active adenylyl cyclase required for meiotic arrest. Dev Biol

71. Oh JS, Han SJ, Conti M. Wee1B, Myt1, and Cdc25 function in distinct compartments of the mouse oocyte to control meiotic resumption. J Cell Biol. 2010;188(2):199–207. 72. Pirino G, Wescott MP, Donovan PJ. Protein kinase a regulates resumption of meiosis by phosphorylation of Cdc25B in mammalian oocytes. Cell Cycle. 2009;8(4):665–670. 73. Masciarelli S, Horner K, Liu C, et al. Cyclic nucleotide phosphodiesterase 3A-­deficient mice as a model of female infertility. J Clin Invest. 2004;114(2):196–205. 74. Robinson JW, Zhang M, Shuhaibar LC, et al. Luteinizing hormone reduces the activity of the NPR2 guanylyl cyclase in mouse ovarian follicles, contributing to the cyclic GMP decrease that promotes resumption of meiosis in oocytes. Dev Biol. 2012;366(2):308–316. 75. Egbert JR, Shuhaibar LC, Edmund AB, et al. Dephosphorylation and inactivation of NPR2 guanylyl cyclase in granulosa cells contributes to the LH-­induced decrease in cGMP that causes resumption of meiosis in rat oocytes. Development. 2014;141(18):3594–3604. 76. Shuhaibar LC, Egbert JR, Norris RP, et al. Intercellular signaling via cyclic GMP diffusion through gap junctions restarts meiosis in mouse ovarian follicles. Proc Natl Acad Sci U S A. 2015;112(17):5527–5532. 77. Mehlmann LM, Saeki Y, Tanaka S, et al. The Gs-­linked receptor GPR3 maintains meiotic arrest in mammalian oocytes. Science. 2004;306(5703):1947–1950. 78. Lincoln AJ, Wickramasinghe D, Stein P, et al. Cdc25b phosphatase is required for resumption of meiosis during oocyte maturation. Nat Genet. 2002;30(4):446–449. 79. Ledan E, Polanski Z, Terret ME, Maro B. Meiotic maturation of the mouse oocyte requires an equilibrium between cyclin B synthesis and degradation. Dev Biol. 2001;232(2):400–413. 80. Reis A, Chang HY, Levasseur M, Jones KT. APCcdh1 activity in mouse oocytes prevents entry into the first meiotic division. Nat Cell Biol. 2006;8(5):539–540. 81. Schindler K, Schultz RM. CDC14B acts through FZR1 (CDH1) to prevent meiotic maturation of mouse oocytes. Biol Reprod. 2009;80(4):795–803. 82. Clift D, Schuh M. A three-­step MTOC fragmentation mechanism facilitates bipolar spindle assembly in mouse oocytes. Nat Commun. 2015;6(1):7217. 83. Holubcová Z, Blayney M, Elder K, Schuh M. Human oocytes. Error-­ prone chromosome-­ mediated spindle assembly favors chromosome segregation defects in human oocytes. Science. 2015;348(6239):1143–1147. 84. Roeles J, Tsiavaliaris G. Actin-­microtubule interplay coordinates spindle assembly in human oocytes. Nat Commun. 2019;10(1):4651. 85. Little TM, Jordan PW. PLK1 is required for chromosome compaction and microtubule organization in mouse oocytes. Mol Biol Cell. 2020;31(12):1206–1217. 86. Blengini CS, Ibrahimian P, Vaskovicova M, Drutovic D, Solc P, Schindler K. Aurora Kinase A Is Essential for Meiosis in Mouse Oocytes. bioRxiv; 2021. Published online. 87. Davydenko O, Schultz RM, Lampson MA. Increased CDK1 activity determines the timing of kinetochore-­microtubule attachments in meiosis I. J Cell Biol. 2013;202(2):221–229. 88. Vallardi G, Cordeiro MH, Saurin AT. A kinase-­phosphatase network that regulates kinetochore-­microtubule attachments and the SAC. Prog Mol Subcell Biol. 2017;56:457–484. 89. Marston AL, Wassmann K. Multiple duties for spindle assembly checkpoint kinases in meiosis. Front Cell Dev Biol. 2017;5:109. 90. Hardwick KG, Weiss E, Luca FC, Winey M, Murray AW. Activation of the budding yeast spindle assembly checkpoint without mitotic spindle disruption. Science. 1996;273(5277):953–956. 91. Hached K, Xie SZ, Buffin E, et al. Mps1 at kinetochores is essential for female mouse meiosis I. Development. 2011;138(11):2261–2271. 92. Brunet S, Maria AS, Guillaud P, Dujardin D, Kubiak JZ, Maro B. Kinetochore fibers are not involved in the formation of the first meiotic spindle in mouse oocytes, but control the exit from the first meiotic M phase. J Cell Biol. 1999;146(1):1–12. 93. Balboula AZ, Schindler K. Selective disruption of aurora C kinase reveals distinct functions from aurora B kinase during meiosis in mouse oocytes. PLoS Genet. 2014;10(2):e1004194. 94. Lane SIR, Yun Y, Jones KT. Timing of anaphase-­ promoting complex activation in mouse oocytes is predicted by microtubule-­ kinetochore attachment but not by bivalent alignment or tension.

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development 95. Sebestova J, Danylevska A, Novakova L, Kubelka M, Anger M. Lack of response to unaligned chromosomes in mammalian female gametes. Cell Cycle. 2012;11(16):3011–3018. 96. Woods LM, Hodges CA, Baart E, Baker SM, Liskay M, Hunt PA. Chromosomal influence on meiotic spindle assembly: abnormal meiosis I in female Mlh1 mutant mice. J Cell Biol. 1999;145(7):1395–1406. 97. LeMaire-­Adkins R, Radke K, Hunt PA. Lack of checkpoint control at the metaphase/anaphase transition: a mechanism of meiotic nondisjunction in mammalian females. J Cell Biol. 1997;139(7):1611–1619. 98. Hunt P, LeMaire R, Embury P, Sheean L, Mroz K. Analysis of chromosome behavior in intact mammalian oocytes: monitoring the segregation of a univalent chromosome during female meiosis. Hum Mol Genet. 1995;4(11):2007–2012. 99. Kouznetsova A, Novak I, Jessberger R, Höög C. SYCP2 and SYCP3 are required for cohesin core integrity at diplotene but not for centromere cohesion at the first meiotic division. J Cell Sci. 2005;118(Pt 10):2271–2278. 100. Pan H, Ma P, Zhu W, Schultz RM. Age-­associated increase in aneuploidy and changes in gene expression in mouse eggs. Dev Biol. 2008;316(2):397–407. 101. Steuerwald N, Cohen J, Herrera RJ, Sandalinas M, Brenner CA. Association between spindle assembly checkpoint expression and maternal age in human oocytes. Mol Hum Reprod. 2001;7(1):49–55. 102. Ottolini CS, Newnham L, Capalbo A, et al. Genome-­wide maps of recombination and chromosome segregation in human oocytes and embryos show selection for maternal recombination rates. Nat Genet. 2015;47(7):727–735. 103. Gruhn JR, Zielinska AP, Shukla V, et al. Chromosome errors in human eggs shape natural fertility over reproductive life span. Science. 2019;365(6460):1466–1469. 104. Lee J, Kitajima TS, Tanno Y, et al. Unified mode of centromeric protection by shugoshin in mammalian oocytes and somatic cells. Nat Cell Biol. 2008;10(1):42–52. 105. Zielinska AP, Holubcova Z, Blayney M, Elder K, Schuh M. Sister kinetochore splitting and precocious disintegration of bivalents could explain the maternal age effect. Elife. 2015;4:e11389. 106. Tachibana-­Konwalski K, Godwin J, van der Weyden L, et al. Rec8-­ containing cohesin maintains bivalents without turnover during the growing phase of mouse oocytes. Genes Dev. 2010;24(22):2505–2516. 107. Burkhardt S, Borsos M, Szydlowska A, et al. Chromosome cohesion established by Rec8-­cohesin in fetal oocytes is maintained without detectable turnover in oocytes arrested for months in mice. Curr Biol. 2016;26(5):678–685. 108. Chiang T, Duncan FE, Schindler K, Schultz RM, Lampson MA. Evidence that weakened centromere cohesion is a leading cause of age-­related aneuploidy in oocytes. Curr Biol. 2010;20(17):1522–1528. 109. Lister LM, Kouznetsova A, Hyslop LA, et al. Age-­related meiotic segregation errors in mammalian oocytes are preceded by depletion of cohesin and Sgo2. Curr Biol. 2010;20(17):1511–1521. 110. Zielinska AP, Bellou E, Sharma N, et al. Meiotic kinetochores fragment into multiple lobes upon cohesin loss in aging eggs. Curr Biol. 2019;29(22):3749–3765.e7. 111. Shomper M, Lappa C, FitzHarris G. Kinetochore microtubule establishment is defective in oocytes from aged mice. Cell Cycle. 2014;13(7):1171–1179. 112. Duncan FE, Schindler K, Schultz RM, et al. Unscrambling the oocyte and the egg: clarifying terminology of the female gamete in mammals. Mol Hum Reprod. 2020;26(11):797–800. 113. Madgwick S, Hansen DV, Levasseur M, Jackson PK, Jones KT. Mouse Emi2 is required to enter meiosis II by reestablishing cyclin B1 during interkinesis. J Cell Biol. 2006;174(6):791–801. 114. Sako K, Suzuki K, Isoda M, et al. Emi2 mediates meiotic MII arrest by competitively inhibiting the binding of Ube2S to the APC/C. Nat Commun. 2014;5(1):3667. 115. Schmidt A, Rauh NR, Nigg EA, Mayer TU. Cytostatic factor: an activity that puts the cell cycle on hold. J Cell Sci. 2006;119(Pt 7):1213–1218. 116. Shoji S, Yoshida N, Amanai M, et al. Mammalian Emi2 mediates cytostatic arrest and transduces the signal for meiotic exit via Cdc20. EMBO J. 2006;25(4):834–845. 117. Ducibella T, Fissore R. The roles of Ca2+, downstream protein kinases, and oscillatory signaling in regulating fertilization and the activation of development. Dev Biol. 2008;315(2):257–279. 118. Verlhac MH, de Pennart H kinase becomes stably activated at metaphase and is associated

216.e3

with microtubule-­organizing centers during meiotic maturation of mouse oocytes. Dev Biol. 1993;158(2):330–340. 119. Verlhac MH, Kubiak JZ, Weber M, et al. Mos is required for MAP kinase activation and is involved in microtubule organization during meiotic maturation in the mouse. Development. 1996;122(3):815–822. 120. Colledge WH, Carlton MB, Udy GB, Evans MJ. Disruption of c-­mos causes parthenogenetic development of unfertilized mouse eggs. Nature. 1994;370(6484):65–68. 121. Chambon JP, Touati SA, Berneau S, et al. The PP2A inhibitor I2PP2A is essential for sister chromatid segregation in oocyte meiosis II. Curr Biol. 2013;23(6):485–490. 122. Sathananthan AH. Ultrastructural changes during meiotic maturation in mammalian oocytes: unique aspects of the human oocyte. Microsc Res Tech. 1994;27(2):145–164. 123. Trebichalská Z, Kyjovská D, Kloudová S, Otevřel P, Hampl A, Holubcová Z. Cytoplasmic maturation in human oocytes: an ultrastructural study. Biol Reprod. 2021;104(1):106–116. 124. Connors SA, Kanatsu-­ Shinohara M, Schultz RM, Kopf GS. Involvement of the cytoskeleton in the movement of cortical granules during oocyte maturation, and cortical granule anchoring in mouse eggs. Dev Biol. 1998;200(1):103–115. 125. Cheeseman LP, Boulanger J, Bond LM, Schuh M. Two pathways regulate cortical granule translocation to prevent polyspermy in mouse oocytes. Nat Commun. 2016;7(1):13726. 126. Vogt EJ, Tokuhiro K, Guo M, et al. Anchoring cortical granules in the cortex ensures trafficking to the plasma membrane for post-­ fertilization exocytosis. Nat Commun. 2019;10(1):2271. 127. Sathananthan AH, Ng SC, Chia CM, Law HY, Edirisinghe WR, Ratnam SS. The origin and distribution of cortical granules in human oocytes with reference to Golgi, nucleolar, and microfilament activity. Ann N Y Acad Sci. 1985;442:251–264. 128. Eppig JJ. The relationship between cumulus cell-­oocyte coupling, oocyte meiotic maturation, and cumulus expansion. Dev Biol. 1982;89(1):268–272. 129. Yi K, Unruh JR, Deng M, Slaughter BD, Rubinstein B, Li R. Dynamic maintenance of asymmetric meiotic spindle position through Arp2/3-­complex-­driven cytoplasmic streaming in mouse oocytes. Nat Cell Biol. 2011;13(10):1252–1258. 130. Zhang B, Zheng H, Huang B, et al. Allelic reprogramming of the histone modification H3K4me3 in early mammalian development. Nature. 2016;537(7621):553–557. 131. Andreu-­Vieyra CV, Chen R, Agno JE, et al. MLL2 is required in oocytes for bulk histone 3 lysine 4 trimethylation and transcriptional silencing. PLoS Biol. 2010;8(8):e1000453. 132. Sankar A, Lerdrup M, Manaf A, et al. KDM4A regulates the maternal-­ to-­ zygotic transition by protecting broad H3K4me3 domains from H3K9me3 invasion in oocytes. Nat Cell Biol. 2020;22(4): 380–388. 133. Lucifero D, Mann MRW, Bartolomei MS, Trasler JM. Gene-­ specific timing and epigenetic memory in oocyte imprinting. Hum Mol Genet. 2004;13(8):839–849. 134. Kong BY, Duncan FE, Que EL, Kim AM, O’Halloran TV, Woodruff TK. Maternally-­ derived zinc transporters ZIP6 and ZIP10 drive the mammalian oocyte-­to-­egg transition. Mol Hum Reprod. 2014;20(11):1077–1089. 135. Kim AM, Vogt S, O’Halloran TV, Woodruff TK. Zinc availability regulates exit from meiosis in maturing mammalian oocytes. Nat Chem Biol. 2010;6(9):674–681. 136. Suzuki T, Yoshida N, Suzuki E, Okuda E, Perry ACF. Full-­term mouse development by abolishing Zn2+-­dependent metaphase II arrest without Ca2+ release. Development. 2010;137(16):2659–2669. 137. Bernhardt ML, Kim AM, O’Halloran TV, Woodruff TK. Zinc requirement during meiosis I-­ meiosis II transition in mouse oocytes is independent of the MOS-­MAPK pathway. Biol Reprod. 2011;84(3):526–536. 138. Tokuhiro K, Dean J. Glycan-­independent gamete recognition triggers egg zinc sparks and ZP2 cleavage to prevent polyspermy. Dev Cell. 2018;46(5):627–640.e5. 139. Bernhardt ML, Stein P, Carvacho I, et al. TRPM7 and CaV3.2 channels mediate Ca2+ influx required for egg activation at fertilization. Proc Natl Acad Sci U S A. 2018;115(44):E10370–E10378. 140. Bernhardt ML, Zhang Y, Erxleben CF, et al. CaV3.2 T-­type channels mediate Ca2+ entry during oocyte maturation and following fertilization. J Cell Sci. 2015;128(23):4442–4452. . Maturation-­associated increase in IP3 receptor type 1: role in conferring increased IP3

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216.e4 PART I  The Fundamentals of Reproduction sensitivity and Ca2+ oscillatory behavior in mouse eggs. Dev Biol. 2003;254(2):163–171. 142. Gambini A, Stein P, Savy V, et al. Developmentally programmed tankyrase activity upregulates β-­catenin and licenses progression of embryonic genome activation. Dev Cell. 2020;53(5):545–560.e7. 143. Medvedev S, Yang J, Hecht NB, Schultz RM. CDC2A (CDK1)-­ mediated phosphorylation of MSY2 triggers maternal mRNA degradation during mouse oocyte maturation. Dev Biol. 2008;321(1):205–215. 144. Gebauer F, Richter JD. Mouse cytoplasmic polyadenylylation element binding protein: an evolutionarily conserved protein that interacts with the cytoplasmic polyadenylylation elements of c-­mos mRNA. Proc Natl Acad Sci U S A. 1996;93(25):14602–14607. 145. Chen J, Melton C, Suh N, et al. Genome-­wide analysis of translation reveals a critical role for deleted in azoospermia-­like (Dazl) at the oocyte-­to-­zygote transition. Genes Dev. 2011;25(7):755–766. 146. Ma J, Fukuda Y, Schultz RM. Mobilization of dormant Cnot7 mRNA promotes deadenylation of maternal transcripts during mouse oocyte maturation. Biol Reprod. 2015;93(2):48. 147. Horvat F, Fulka H, Jankele R, et al. Role of Cnot6l in maternal mRNA turnover. Life Sci Alliance. 2018;1(4):e201800084. 148. Morgan M, Much C, DiGiacomo M, et al. mRNA 3′ uridylation and poly(A) tail length sculpt the mammalian maternal transcriptome. Nature. 2017;548(7667):347–351. 149. Lim J, Ha M, Chang H, et al. Uridylation by TUT4 and TUT7 marks mRNA for degradation. Cell. 2014;159(6):1365–1376. 150. Ma J, Flemr M, Strnad H, Svoboda P, Schultz RM. Maternally recruited DCP1A and DCP2 contribute to messenger RNA degradation during oocyte maturation and genome activation in mouse. Biol Reprod. 2013;88(1):11. 151. Sha QQ, Zheng W, Wu YW, et al. Dynamics and clinical relevance of maternal mRNA clearance during the oocyte-­to-­embryo transition in humans. Nat Commun. 2020;11(1):4917. 152. Perreault SD, Barbee RR, Slott VL. Importance of glutathione in the acquisition and maintenance of sperm nuclear decondensing activity in maturing hamster oocytes. Dev Biol. 1988;125(1):181–186. 153. Combelles CMH, Cekleniak NA, Racowsky C, Albertini DF. Assessment of nuclear and cytoplasmic maturation in in-­ vitro matured human oocytes. Hum Reprod. 2002;17(4):1006–1016. 154. Kan R, Yurttas P, Kim B, et al. Regulation of mouse oocyte microtubule and organelle dynamics by PADI6 and the cytoplasmic lattices. Dev Biol. 2011;350(2):311–322. 155. Cohen J, McNaughton DC. Spermatozoa: the probable selection of a small population by the genital tract of the female rabbit. J Reprod Fertil. 1974;39(2):297–310. 156. Siddiquey AK, Cohen J. In-­vitro fertilization in the mouse and the relevance of different sperm/egg concentrations and volumes. J Reprod Fertil. 1982;66(1):237–242. 157. Fischer B, Adams CE. Fertilization following mixed insemination with “cervix-­selected” and “unselected” spermatozoa in the rabbit. J Reprod Fertil. 1981;62(2):337–343. 158. Gould JE, Overstreet JW, Hanson FW. Assessment of human sperm function after recovery from the female reproductive tract. Biol Reprod. 1984;31(5):888–894. 159. van der Linden PJ, Kets M, Gimpel JA, Wiegerinck MA. Cyclic changes in the concentration of glucose and fructose in human cervical mucus. Fertil Steril. 1992;57(3):573–577. 160. Settlage DS, Motoshima M, Tredway DR. Sperm transport from the external cervical os to the fallopian tubes in women: a time and quantitation study. Fertil Steril. 1973;24(9):655–661. 161. Kunz G, Beil D, Deininger H, Wildt L, Leyendecker G. The dynamics of rapid sperm transport through the female genital tract: evidence from vaginal sonography of uterine peristalsis and hysterosalpingoscintigraphy. Hum Reprod. 1996;11(3):627–632. 162. Zervomanolakis I, Ott HW, Hadziomerovic D, et al. Physiology of upward transport in the human female genital tract. Ann N Y Acad Sci. 2007;1101(1):1–20. 163. Ishikawa Y, Usui T, Yamashita M, Kanemori Y, Baba T. Surfing and swimming of ejaculated sperm in the mouse oviduct. Biol Reprod. 2016;94(4):89. 164. van Gestel I, IJland MM, Hoogland HJ, Evers JLH. Endometrial wave-­like activity in the non-­pregnant uterus. Hum Reprod Update. 2003;9(2):131–138. 165. Blasco V, Prados N, Carranza F, González-­Ravina C, Pellicer A, Fernández-­Sánchez M. Influence of follicle rupture and uterine contractions on intrauterine insemination outcome: a new predic tive model. Fertil Steril

166. Wilcox AJ, Weinberg CR, Baird DD. Timing of sexual intercourse in relation to ovulation. Effects on the probability of conception, survival of the pregnancy, and sex of the baby. N Engl J Med. 1995;333(23):1517–1521. 167. Kawano N, Araki N, Yoshida K, et al. Seminal vesicle protein SVS2 is required for sperm survival in the uterus. Proc Natl Acad Sci U S A. 2014;111(11):4145–4150. 168. Suarez SS, Pacey AA. Sperm transport in the female reproductive tract. Hum Reprod Update. 2006;12(1):23–37. 169. Gwathmey TM, Ignotz GG, Suarez SS. PDC-­109 (BSP-­A1/A2) promotes bull sperm binding to oviductal epithelium in vitro and may be involved in forming the oviductal sperm reservoir. Biol Reprod. 2003;69(3):809–815. 170. Suarez SS. Mammalian sperm interactions with the female reproductive tract. Cell Tissue Res. 2016;363(1):185–194. 171. Yamaguchi R, Muro Y, Isotani A, et al. Disruption of ADAM3 impairs the migration of sperm into oviduct in mouse. Biol Reprod. 2009;81(1):142–146. 172. Ignotz GG, Cho MY, Suarez SS. Annexins are candidate oviductal receptors for bovine sperm surface proteins and thus may serve to hold bovine sperm in the oviductal reservoir. Biol Reprod. 2007;77(6):906–913. 173. Fujii DT, Yohannes E, Por ED, et al. The proteome of human Fallopian tube lavages during the phase of embryo transit reveals candidate proteins for the optimization of preimplantation embryo culture. Hum Reprod. 2021;36(2):367–380. 174. Dobrinski I, Smith TT, Suarez SS, Ball BA. Membrane contact with oviductal epithelium modulates the intracellular calcium concentration of equine spermatozoa in vitro. Biol Reprod. 1997;56(4):861–869. 175. Ellington JE, Ignotz GG, Varner DD, Marcucio RS, Mathison P, Ball BA. In vitro interaction between oviduct epithelia and equine sperm. Arch Androl. 1993;31(2):79–86. 176. Kawakami E, Kashiwagi C, Hori T, Tsutsui T. Effects of canine oviduct epithelial cells on movement and capacitation of homologous spermatozoa in vitro. Anim Reprod Sci. 2001;68(1–2):121–131. 177. Murray SC, Smith TT. Sperm interaction with fallopian tube apical membrane enhances sperm motility and delays capacitation. Fertil Steril. 1997;68(2):351–357. 178. Smith TT, Yanagimachi R. Attachment and release of spermatozoa from the caudal isthmus of the hamster oviduct. J Reprod Fertil. 1991;91(2):567–573. 179. Ho HC, Suarez SS. Hyperactivation of mammalian spermatozoa: function and regulation. J Reprod Fertil. 2001;122(4):519–526. 180. Ho K, Wolff CA, Suarez SS. CatSper-­null mutant spermatozoa are unable to ascend beyond the oviductal reservoir. Reprod Fertil Dev. 2009;21(2):345–350. 181. Ralt D, Manor M, Cohen-­Dayag A, et al. Chemotaxis and chemokinesis of human spermatozoa to follicular factors. Biol Reprod. 1994;50(4):774–785. 182. Jaiswal BS, Tur-­Kaspa I, Dor J, Mashiach S, Eisenbach M. Human sperm chemotaxis: is progesterone a chemoattractant? Biol Reprod. 1999;60(6):1314–1319. 183. Wachten D, Jikeli JF, Kaupp UB. Sperm sensory signaling. Cold Spring Harb Perspect Biol. 2017;9(7):a028225. 184. Hino T, Yanagimachi R. Active peristaltic movements and fluid production of the mouse oviduct: their roles in fluid and sperm transport and fertilization. Biol Reprod. 2019;101(1):40–49. 185. Yanagimachi R. The movement of golden hamster spermatozoa before and after capacitation. J Reprod Fertil. 1970;23(1):193–196. 186. Demott RP, Suarez SS. Hyperactivated sperm progress in the mouse oviduct. Biol Reprod. 1992;46(5):779–785. 187. Fitzpatrick JL, Willis C, Devigili A, et al. Chemical signals from eggs facilitate cryptic female choice in humans. Proc Biol Sci. 2020;287(1928):20200805. 188. Ralt D, Goldenberg M, Fetterolf P, et al. Sperm attraction to a follicular factor(s) correlates with human egg fertilizability. Obstet Gynecol Surv. 1991;46(9):648. 189. Bahat A, Caplan SR, Eisenbach M. Thermotaxis of human sperm cells in extraordinarily shallow temperature gradients over a wide range. PLoS One. 2012;7(7):e41915. 190. Bahat A, Tur-­Kaspa I, Gakamsky A, Giojalas LC, Breitbart H, Eisenbach M. Thermotaxis of mammalian sperm cells: a potential navigation mechanism in the female genital tract. Nat Med. 2003;9(2):149–150. . Observations on the penetration of the sperm into the mammalian egg. J

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development 192. Chang MC. Fertilizing capacity of spermatozoa deposited into the fallopian tubes. Nature. 1951;168(4277):697–698. 193. Austin CR. The capacitation of the mammalian sperm. Nature. 1952;170(4321):326. 194. Yanagimachi R, Chang MC. Fertilization of hamster eggs in vitro. Nature. 1963;200(4903):281–282. 195. Chang MC. A detrimental effect of seminal plasma on the fertilizing capacity of sperm. Nature. 1957;179(4553):258–259. 196. Davis BK. Decapacitation and recapacitation of rabbit spermatozoa treated with membrane vesicles from seminal plasma. J Reprod Fertil. 1974;41(1):241–244. 197. Dukelow WR, Chernoff HN, Williams WL. Properties of decapacitation factor and presence in various species. J Reprod Fertil. 1967;14(3):393–399. 198. Davis BK. Timing of fertilization in mammals: sperm cholesterol/ phospholipid ratio as a determinant of the capacitation interval. Proc Natl Acad Sci U S A. 1981;78(12):7560–7564. 199. Kawano N, Yoshida M. Semen-­ coagulating protein, SVS2, in mouse seminal plasma controls sperm fertility. Biol Reprod. 2007;76(3):353–361. 200. Kawano N, Yoshida K, Iwamoto T, Yoshida M. Ganglioside GM1 mediates decapacitation effects of SVS2 on murine spermatozoa. Biol Reprod. 2008;79(6):1153–1159. 201. Nixon B, MacIntyre DA, Mitchell LA, Gibbs GM, O’Bryan M, Aitken RJ. The identification of mouse sperm-­surface-­associated proteins and characterization of their ability to act as decapacitation factors. Biol Reprod. 2006;74(2):275–287. 202. Robert M, Gagnon C. Purification and characterization of the active precursor of a human sperm motility inhibitor secreted by the seminal vesicles: identity with semenogelin. Biol Reprod. 1996;55(4):813–821. 203. de Lamirande E, Yoshida K, Yoshiike TM, Iwamoto T, Semenogelin GC. The main protein of semen coagulum, inhibits human sperm capacitation by interfering with the superoxide anion generated during this process. J Androl. 2001;22(4):672–679. 204. Cross NL. Human seminal plasma prevents sperm from becoming acrosomally responsive to the agonist, progesterone: cholesterol is the major inhibitor. Biol Reprod. 1996;54(1):138–145. 205. Cross NL. Decrease in order of human sperm lipids during capacitation. Biol Reprod. 2003;69(2):529–534. 206. Buck J, Sinclair ML, Schapal L, Cann MJ, Levin LR. Cytosolic adenylyl cyclase defines a unique signaling molecule in mammals. Proc Natl Acad Sci U S A. 1999;96(1):79–84. 207. Chen Y, Cann MJ, Litvin TN, et al. Soluble adenylyl cyclase as an evolutionarily conserved bicarbonate sensor. Science. 2000;289(5479):625–628. 208. Esposito G, Jaiswal BS, Xie F, et al. Mice deficient for soluble adenylyl cyclase are infertile because of a severe sperm-­motility defect. Proc Natl Acad Sci U S A. 2004;101(9):2993–2998. 209. Visconti PE, Moore GD, Bailey JL, et al. Capacitation of mouse spermatozoa. II. Protein tyrosine phosphorylation and capacitation are regulated by a cAMP-­dependent pathway. Development. 1995;121(4):1139–1150. 210. Visconti PE, Bailey JL, Moore GD, Pan D, Olds-­Clarke P, Kopf GS. Capacitation of mouse spermatozoa. 1. Correlation between the capacitation state and protein tyrosine phosphorylation. Development. 1995;121:1129–1137. 211. Osheroff JE, Visconti PE, Valenzuela JP, Travis AJ, Alvarez J, Kopf GS. Regulation of human sperm capacitation by a cholesterol efflux-­ stimulated signal transduction pathway leading to protein kinase A-­mediated up-­regulation of protein tyrosine phosphorylation. Mol Hum Reprod. 1999;5(11):1017–1026. 212. Gadella BM. Sperm membrane physiology and relevance for fertilization. Anim Reprod Sci. 2008;107(3–4):229–236. 213. Travis AJ, Kopf GS. The role of cholesterol efflux in regulating the fertilization potential of mammalian spermatozoa. J Clin Invest. 2002;110(6):731–736. 214. Cross NL. Role of cholesterol in sperm capacitation. Biol Reprod. 1998;59(1):7–11. 215. Lishko PV, Botchkina IL, Fedorenko A, Kirichok Y. Acid extrusion from human spermatozoa is mediated by flagellar voltage-­gated proton channel. Cell. 2010;140(3):327–337. 216. Puga Molina LC, Luque GM, Balestrini PA, Marín-­Briggiler CI, Romarowski A, Buffone MG. Molecular basis of human sperm capacitation. Front

216.e5

217. Wang D, King SM, Quill TA, Doolittle LK, Garbers DL. A new sperm-­specific Na+/H+ exchanger required for sperm motility and fertility. Nat Cell Biol. 2003;5(12):1117–1122. 218. Puga Molina LC, Pinto NA, Torres Rodríguez P, et al. Essential role of CFTR in PKA-­dependent phosphorylation, alkalinization, and hyperpolarization during human sperm capacitation. J Cell Physiol. 2017;232(6):1404–1414. 219. Xu WM, Shi QX, Chen WY, et al. Cystic fibrosis transmembrane conductance regulator is vital to sperm fertilizing capacity and male fertility. Proc Natl Acad Sci U S A. 2007;104(23):9816–9821. 220. Li CY, Jiang LY, Chen WY, et al. CFTR is essential for sperm fertilizing capacity and is correlated with sperm quality in humans. Hum Reprod. 2010;25(2):317–327. 221. Wedenoja S, Khamaysi A, Shimshilashvili L, et al. A missense mutation in SLC26A3 is associated with human male subfertility and impaired activation of CFTR. Sci Rep. 2017;7(1):14208. 222. Zeng Y, Clark EN, Florman HM. Sperm membrane potential: hyperpolarization during capacitation regulates zona pellucida-­ dependent acrosomal secretion. Dev Biol. 1995;171(2):554–563. 223. Mannowetz N, Naidoo NM, Choo SAS, Smith JF, Lishko PV. Slo1 is the principal potassium channel of human spermatozoa. Elife. 2013;2:e01009. 224. Santi CM, Martínez-­López P, de la Vega-­Beltrán JL, et al. The SLO3 sperm-­specific potassium channel plays a vital role in male fertility. FEBS Lett. 2010;584(5):1041–1046. 225. López-­González I, Torres-­Rodríguez P, Sánchez-­Carranza O, et al. Membrane hyperpolarization during human sperm capacitation. Mol Hum Reprod. 2014;20(7):619–629. 226. McDermott J, Sánchez G, Nangia AK, Blanco G. Role of human Na,K-­ATPase alpha 4 in sperm function, derived from studies in transgenic mice. Mol Reprod Dev. 2015;82(3):167–181. 227. Puga Molina LC, Pinto NA, Torres NI, et al. CFTR/ENaC-­ dependent regulation of membrane potential during human sperm capacitation is initiated by bicarbonate uptake through NBC. J Biol Chem. 2018;293(25):9924–9936. 228. De Blas G, Michaut M, Treviño CL, et al. The intraacrosomal calcium pool plays a direct role in acrosomal exocytosis. J Biol Chem. 2002;277(51):49326–49331. 229. Herrick SB, Schweissinger DL, Kim SW, Bayan KR, Mann S, Cardullo RA. The acrosomal vesicle of mouse sperm is a calcium store. J Cell Physiol. 2005;202(3):663–671. 230. Ho HC, Suarez SS. Characterization of the intracellular calcium store at the base of the sperm flagellum that regulates hyperactivated motility. Biol Reprod. 2003;68(5):1590–1596. 231. Schuh K, Cartwright EJ, Jankevics E, et al. Plasma membrane Ca2+ ATPase 4 is required for sperm motility and male fertility. J Biol Chem. 2004;279(27):28220–28226. 232. Krasznai Z, Krasznai ZT, Morisawa M, et al. Role of the Na+/Ca2+ exchanger in calcium homeostasis and human sperm motility regulation. Cell Motil Cytoskeleton. 2006;63(2):66–76. 233. Sakata Y, Saegusa H, Zong S, et al. Cav2.3 (α1E) Ca2+ channel participates in the control of sperm function. FEBS Lett. 2002;516(1–3):229–233. 234. Ren D, Navarro B, Perez G, et al. A sperm ion channel required for sperm motility and male fertility. Nature. 2001;413(6856):603–609. 235. Cohen R, Buttke DE, Asano A, et al. Lipid modulation of calcium flux through CaV2.3 regulates acrosome exocytosis and fertilization. Dev Cell. 2014;28(3):310–321. 236. Avenarius MR, Hildebrand MS, Zhang Y, et al. Human male infertility caused by mutations in the CATSPER1 channel protein. Am J Hum Genet. 2009;84(4):505–510. 237. Brown SG, Miller MR, Lishko PV, et al. Homozygous in-­frame deletion in CATSPERE in a man producing spermatozoa with loss of CatSper function and compromised fertilizing capacity. Hum Reprod. 2018;33(10):1812–1816. 238. Smith JF, Syritsyna O, Fellous M, et al. Disruption of the principal, progesterone-­activated sperm Ca2+ channel in a CatSper2-­deficient infertile patient. Proc Natl Acad Sci U S A. 2013;110(17):6823–6828. 239. Williams HL, Mansell S, Alasmari W, et al. Specific loss of CatSper function is sufficient to compromise fertilizing capacity of human spermatozoa. Hum Reprod. 2015;30(12):2737–2746. 240. Hwang JY, Mannowetz N, Zhang Y, et al. Dual sensing of physiologic pH and calcium by EFCAB9 regulates sperm motility. Cell. 2019;177(6):1480–1494.e19.

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216.e6 PART I  The Fundamentals of Reproduction 241. Rahban R, Nef S. CatSper: the complex main gate of calcium entry in mammalian spermatozoa. Mol Cell Endocrinol. 2020;518(110951):110951. 242. Chung JJ, Shim SH, Everley RA, Gygi SP, Zhuang X, Clapham DE. Structurally distinct Ca(2+) signaling domains of sperm flagella orchestrate tyrosine phosphorylation and motility. Cell. 2014;157(4):808–822. 243. Lishko PV, Botchkina IL, Kirichok Y. Progesterone activates the principal Ca2+ channel of human sperm. Nature. 2011;471(7338):387–391. 244. Orta G, de la Vega-­ Beltran JL, Martín-­ Hidalgo D, Santi CM, Visconti PE, Darszon A. CatSper channels are regulated by protein kinase A. J Biol Chem. 2018;293(43):16830–16841. 245. Kirichok Y, Navarro B, Clapham DE. Whole-­cell patch-­clamp measurements of spermatozoa reveal an alkaline-­ activated Ca2+ channel. Nature. 2006;439(7077):737–740. 246. Miller MR, Mannowetz N, Iavarone AT, et al. Unconventional endocannabinoid signaling governs sperm activation via the sex hormone progesterone. Science. 2016;352(6285):555–559. 247. Lin Y, Mahan K, Lathrop WF, Myles DG, Primakoff P. A hyaluronidase activity of the sperm plasma membrane protein PH-­20 enables sperm to penetrate the cumulus cell layer surrounding the egg. J Cell Biol. 1994;125(5):1157–1163. 248. Kim E, Yamashita M, Kimura M, Honda A, Kashiwabara SI, Baba T. Sperm penetration through cumulus mass and zona pellucida. Int J Dev Biol. 2008;52(5–6):677–682. 249. Lefièvre L, Conner SJ, Salpekar A, et al. Four zona pellucida glycoproteins are expressed in the human. Hum Reprod. 2004;19(7):1580–1586. 250. Baibakov B, Boggs NA, Yauger B, Baibakov G, Dean J. Human sperm bind to the N-­terminal domain of ZP2 in humanized zonae pellucidae in transgenic mice. J Cell Biol. 2012;197(7):897–905. 251. Avella MA, Baibakov B, Dean J. A single domain of the ZP2 zona pellucida protein mediates gamete recognition in mice and humans. J Cell Biol. 2014;205(6):801–809. 252. Bhakta HH, Refai FH, Avella MA. The molecular mechanisms mediating mammalian fertilization. Development. 2019;146(15): dev176966. 253. Tardif S, Wilson MD, Wagner R, et al. Zonadhesin is essential for species specificity of sperm adhesion to the egg zona pellucida. J Biol Chem. 2010;285(32):24863–24870. 254. Ensslin MA, Shur BD. Identification of mouse sperm SED1, a bimotif EGF repeat and discoidin-­domain protein involved in sperm-­egg binding. Cell. 2003;114(4):405–417. 255. Muro Y, Buffone MG, Okabe M, Gerton GL. Function of the acrosomal matrix: zona pellucida 3 receptor (ZP3R/sp56) is not essential for mouse fertilization. Biol Reprod. 2012;86(1):1–6. 256. Lu Q, Shur BD. Sperm from beta 1,4-­galactosyltransferase-­null mice are refractory to ZP3-­induced acrosome reactions and penetrate the zona pellucida poorly. Development. 1997;124(20):4121–4131. 257. Fleming AD, Yanagimachi R. Fertile life of acrosome-­ reacted Guinea pig spermatozoa. J Exp Zool. 1982;220(1):109–115. 258. Inoue N, Ikawa M, Isotani A, Okabe M. The immunoglobulin superfamily protein Izumo is required for sperm to fuse with eggs. Nature. 2005;434(7030):234–238. 259. Toshimori K. Penetration of the mouse sperm head through the zona pellucida in vivo: an electronmicroscope study at 200KV. Biol Reprod. 1982;26(3):475–481. 260. Kuzan FB, Fleming AD, Seidel Jr GE. Successful fertilization in vitro of fresh intact oocytes by perivitelline (acrosome-­reacted) spermatozoa of the rabbit. Fertil Steril. 1984;41(5):766–770. 261. Muro Y, Hasuwa H, Isotani A, et al. Behavior of mouse spermatozoa in the female reproductive tract from soon after mating to the beginning of fertilization. Biol Reprod. 2016;94(4):80. 262. La Spina FA, Puga Molina LC, Romarowski A, et al. Mouse sperm begin to undergo acrosomal exocytosis in the upper isthmus of the oviduct. Dev Biol. 2016;411(2):172–182. 263. Hino T, Muro Y, Tamura-­ Nakano M, Okabe M, Tateno H, Yanagimachi R. The behavior and acrosomal status of mouse spermatozoa in vitro, and within the oviduct during fertilization after natural mating. Biol Reprod. 2016;95(3):50. 264. Inoue N, Satouh Y, Ikawa M, Okabe M, Yanagimachi R. Acrosome-­ reacted mouse spermatozoa recovered from the perivitelline space can fertilize other eggs. Proc Natl Acad Sci U S A. 2011;108(50):20008–20011.

265. Baba T, Azuma S, Kashiwabara S, Toyoda Y. Sperm from mice carrying a targeted mutation of the acrosin gene can penetrate the oocyte zona pellucida and effect fertilization. J Biol Chem. 1994;269(50): 31845–31849. 266. Yamashita M, Honda A, Ogura A, Kashiwabara SI, Fukami K, Baba T. Reduced fertility of mouse epididymal sperm lacking Prss21/ Tesp5 is rescued by sperm exposure to uterine microenvironment. Gene Cell. 2008;13(10):1001–1013. 267. Kawano N, Kang W, Yamashita M, et al. Mice lacking two sperm serine proteases, ACR and PRSS21, are subfertile, but the mutant sperm are infertile in vitro. Biol Reprod. 2010;83(3):359–369. 268. Adham IM, Nayernia K, Engel W. Spermatozoa lacking acrosin protein show delayed fertilization. Mol Reprod Dev. 1997;46(3):370–376. 269. Hirose M, Honda A, Fulka H, et al. Acrosin is essential for sperm penetration through the zona pellucida in hamsters. Proc Natl Acad Sci U S A. 2020;117(5):2513–2518. 270. Shamsadin R, Adham IM, Nayernia K, Heinlein UA, Oberwinkler H, Engel W. Male mice deficient for germ-­cell cyritestin are infertile. Biol Reprod. 1999;61(6):1445–1451. 271. Nishimura H, Kim E, Nakanishi T, Baba T. Possible function of the ADAM1a/ADAM2 Fertilin complex in the appearance of ADAM3 on the sperm surface. J Biol Chem. 2004;279(33):34957–34962. 272. Krege JH, John SW, Langenbach LL, et al. Male-­female differences in fertility and blood pressure in ACE-­deficient mice. Nature. 1995;375(6527):146–148. 273. Ikawa M, Wada I, Kominami K, et al. The putative chaperone calmegin is required for sperm fertility. Nature. 1997;387(6633):607–611. 274. Yamaguchi R, Yamagata K, Ikawa M, Moss SB, Okabe M. Aberrant distribution of ADAM3 in sperm from both angiotensin-­converting enzyme (Ace)-­and calmegin (Clgn)-­ deficient mice. Biol Reprod. 2006;75(5):760–766. 275. Bedford JM, Moore HD, Franklin LE. Significance of the equatorial segment of the acrosome of the spermatozoon in eutherian mammals. Exp Cell Res. 1979;119(1):119–126. 276. Bianchi E, Doe B, Goulding D, Wright GJ. Juno is the egg Izumo receptor and is essential for mammalian fertilization. Nature. 2014;508(7497):483–487. 277. Satouh Y, Inoue N, Ikawa M, Okabe M. Visualization of the moment of mouse sperm-­egg fusion and dynamic localization of IZUMO1. J Cell Sci. 2012;125(Pt 21):4985–4990. 278. Inoue N, Hamada D, Kamikubo H, et al. Molecular dissection of IZUMO1, a sperm protein essential for sperm-­ egg fusion. Development. 2013;140(15):3221–3229. 279. Aydin H, Sultana A, Li S, Thavalingam A, Lee JE. Molecular architecture of the human sperm IZUMO1 and egg JUNO fertilization complex. Nature. 2016;534(7608):562–565. 280. Ohto U, Ishida H, Krayukhina E, Uchiyama S, Inoue N, Shimizu T. Structure of IZUMO1-­JUNO reveals sperm-­oocyte recognition during mammalian fertilization. Nature. 2016;534(7608):566–569. 281. Noda T, Lu Y, Fujihara Y, et al. Sperm proteins SOF1, TMEM95, and SPACA6 are required for sperm-­oocyte fusion in mice. Proc Natl Acad Sci U S A. 2020;117(21):11493–11502. 282. Fujihara Y, Lu Y, Noda T, et al. Spermatozoa lacking Fertilization Influencing Membrane Protein (FIMP) fail to fuse with oocytes in mice. Proc Natl Acad Sci U S A. 2020;117(17):9393–9400. 283. Lorenzetti D, Poirier C, Zhao M, Overbeek PA, Harrison W, Bishop CE. A transgenic insertion on mouse chromosome 17 inactivates a novel immunoglobulin superfamily gene potentially involved in sperm-­egg fusion. Mamm Genome. 2014;25(3–4):141–148. 284. Lamas-­Toranzo I, Hamze JG, Bianchi E, et al. TMEM95 is a sperm membrane protein essential for mammalian fertilization. Elife. 2020;9:e53913. 285. Pausch H, Kölle S, Wurmser C, et al. A nonsense mutation in TMEM95 encoding a nondescript transmembrane protein causes idiopathic male subfertility in cattle. PLoS Genet. 2014;10(1):e1004044. 286. Fernandez-­Fuertes B, Laguna-­Barraza R, Fernandez-­Gonzalez R, et al. Subfertility in bulls carrying a nonsense mutation in transmembrane protein 95 is due to failure to interact with the oocyte vestments. Biol Reprod. 2017;97(1):50–60. 287. Ellerman DA, Pei J, Gupta S, Snell WJ, Myles D, Primakoff P. Izumo is part of a multiprotein family whose members form large complexes on mammalian sperm: characterization of IZUMO fam. 2009;76(12):1188–1199.

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development 288. Le Naour F, Rubinstein E, Jasmin C, Prenant M, Boucheix C. Severely reduced female fertility in CD9-­deficient mice. Science. 2000;287(5451):319–321. 289. Miyado K, Yamada G, Yamada S, et al. Requirement of CD9 on the egg plasma membrane for fertilization. Science. 2000;287(5451):321–324. 290. Kaji K, Oda S, Shikano T, et al. The gamete fusion process is defective in eggs of Cd9-­deficient mice. Nat Genet. 2000;24(3):279–282. 291. Rubinstein E, Ziyyat A, Prenant M, et al. Reduced fertility of female mice lacking CD81. Dev Biol. 2006;290(2):351–358. 292. Kaji K, Oda S, Miyazaki S, Kudo A. Infertility of CD9-­deficient mouse eggs is reversed by mouse CD9, human CD9, or mouse CD81; polyadenylated mRNA injection developed for molecular analysis of sperm-­egg fusion. Dev Biol. 2002;247(2):327–334. 293. Runge KE, Evans JE, He ZY, et al. Oocyte CD9 is enriched on the microvillar membrane and required for normal microvillar shape and distribution. Dev Biol. 2007;304(1):317–325. 294. Almeida EAC, Huovila APJ, Sutherland AE, et al. Mouse egg integrin α6β1functions as a sperm receptor. Cell. 1995;81(7):1095–1104. 295. Miller BJ, Georges-­Labouesse E, Primakoff P, Myles DG. Normal fertilization occurs with eggs lacking the integrin alpha6beta1 and is CD9-­dependent. J Cell Biol. 2000;149(6):1289–1296. 296. He ZY, Brakebusch C, Fässler R, Kreidberg JA, Primakoff P, Myles DG. None of the integrins known to be present on the mouse egg or to be ADAM receptors are essential for sperm-­egg binding and fusion. Dev Biol. 2003;254(2):226–237. 297. Tsuiki A, Hoshiai H, Takahashi K, Suzuki M, Hoshi K. Sperm-­egg interactions observed by scanning electron microscopy. Arch Androl. 1986;16(1):35–47. 298. Phillips DM, Shalgi R. Sperm penetration into rat ova fertilized in vivo. J Exp Zool. 1982;221(3):373–378. 299. Yanagimachi R, Noda YD. Electron microscope studies of sperm incorporation into the golden hamster egg. Am J Anat. 1970;128(4):429–462. 300. Courtot AM, Feinberg JM, Schoevaert DA, Rainteau DP, Weinman SJ. Calmodulin during human sperm incorporation into hamster oocyte: an immunogold electron microscope study. Mol Reprod Dev. 1994;38(2):170–177. 301. Ravaux B, Garroum N, Perez E, Willaime H, Gourier C. A specific flagellum beating mode for inducing fusion in mammalian fertilization and kinetics of sperm internalization. Sci Rep. 2016;6:31886. 302. Kumakiri J, Oda S, Kinoshita K, Miyazaki S. Involvement of Rho family G protein in the cell signaling for sperm incorporation during fertilization of mouse eggs: inhibition by clostridium difficile toxin B. Dev Biol. 2003;260(2):522–535. 303. Sutovsky P, Navara CS, Schatten G. Fate of the sperm mitochondria, and the incorporation, conversion, and disassembly of the sperm tail structures during bovine fertilization. Biol Reprod. 1996;55(6):1195–1205. 304. Rogers BJ, Bastias C, Coulson RL, Russell LD. Cytochalasin D inhibits penetration of hamster eggs by Guinea pig and human spermatozoa. J Androl. 1989;10(4):275–282. 305. Sun QY, Lai L, Park KW, Kühholzer B, Prather RS, Schatten H. Dynamic events are differently mediated by microfilaments, microtubules, and mitogen-­activated protein kinase during porcine oocyte maturation and fertilization in vitro. Biol Reprod. 2001;64(3):879–889. 306. Perreault SD, Wolff RA, Zirkin BR. The role of disulfide bond reduction during mammalian sperm nuclear decondensation in vivo. Dev Biol. 1984;101(1):160–167. 307. Poccia D. Remodeling of nucleoproteins during gametogenesis, fertilization, and early development. Int Rev Cytol. 1986;105:1–65. 308. Gou LT, Lim DH, Ma W, et al. Initiation of parental genome reprogramming in fertilized oocyte by splicing kinase SRPK1-­catalyzed protamine phosphorylation. Cell. 2020;180(6):1212–1227.e14. 309. Inoue A, Ogushi S, Saitou M, Suzuki MG, Aoki F. Involvement of mouse nucleoplasmin 2 in the decondensation of sperm chromatin after fertilization. Biol Reprod. 2011;85(1):70–77. 310. Stein P, Savy V, Williams AM, Williams CJ. Modulators of calcium signalling at fertilization. Open Biol. 2020;10(7):200118. 311. Saunders CM, Larman MG, Parrington J, et al. PLCζ: a sperm-­ specific trigger of Ca2+oscillations in eggs and embryo development. Development. 2002;129:3533–3544. 312. Heytens E, Parrington J, Coward K, et al. Reduced amounts and abnormal forms of phospholipase C zeta (PLCzeta) in spermatozoa from infertile men.

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313. Kashir J, Konstantinidis M, Jones C, et al. A maternally inherited autosomal point mutation in human phospholipase C zeta (PLCζ) leads to male infertility. Hum Reprod. 2012;27(1):222–231. 314. Torra-­Massana M, Cornet-­Bartolomé D, Barragán M, et al. Novel phospholipase C zeta 1 mutations associated with fertilization failures after ICSI. Hum Reprod. 2019;34(8):1494–1504. 315. Dai J, Dai C, Guo J, et al. Novel homozygous variations in PLCZ1 lead to poor or failed fertilization characterized by abnormal localization patterns of PLCζ in sperm. Clin Genet. 2020;97(2):347–351. 316. Ferrer-­Vaquer A, Barragan M, Freour T, Vernaeve V, Vassena R. PLCζ sequence, protein levels, and distribution in human sperm do not correlate with semen characteristics and fertilization rates after ICSI. J Assist Reprod Genet. 2016;33(6):747–756. 317. Escoffier J, Lee HC, Yassine S, et al. Homozygous mutation of PLCZ1 leads to defective human oocyte activation and infertility that is not rescued by the WW-­binding protein PAWP. Hum Mol Genet. 2016;25(5):878–891. 318. Miyazaki S, Yuzaki M, Nakada K, et al. Block of Ca2+ wave and Ca2+ oscillation by antibody to the inositol 1,4,5-­trisphosphate receptor in fertilized hamster eggs. Science. 1992;257(5067):251–255. 319. Xu Z, Kopf GS, Schultz RM. Involvement of inositol 1,4,5-­trisphosphate-­mediated Ca2+ release in early and late events of mouse egg activation. Development. 1994;120(7):1851–1859. 320. Dumollard R, Marangos P, Fitzharris G, Swann K, Duchen M, Carroll J. Sperm-­triggered [Ca2+] oscillations and Ca2+ homeostasis in the mouse egg have an absolute requirement for mitochondrial ATP production. Development. 2004;131(13):3057–3067. 321. Campbell K, Swann K. Ca2+ oscillations stimulate an ATP increase during fertilization of mouse eggs. Dev Biol. 2006;298(1):225–233. 322. Bernhardt ML. TRPM7 and CaV3.2 channels mediate Ca2+ influx required for egg activation at fertilization. Proc Natl Acad Sci U S A. 2018;115:E10370–E10378. 323. Carvacho I, Lee HC, Fissore RA, Clapham DE. TRPV3 channels mediate strontium-­ induced mouse-­ egg activation. Cell Rep. 2013;5(5):1375–1386. 324. Ozil JP, Markoulaki S, Toth S, et al. Egg activation events are regulated by the duration of a sustained [Ca2+]cyt signal in the mouse. Dev Biol. 2005;282(1):39–54. 325. Ozil JP, Banrezes B, Tóth S, Pan H, Schultz RM. Ca2+ oscillatory pattern in fertilized mouse eggs affects gene expression and development to term. Dev Biol. 2006;300(2):534–544. 326. Tóth S, Huneau D, Banrezes B, Ozil JP. Egg activation is the result of calcium signal summation in the mouse. J Reprod Fertil. 2006;131(1):27–34. 327. Hudmon A, Schulman H. Neuronal CA2+/calmodulin-­dependent protein kinase II: the role of structure and autoregulation in cellular function. Annu Rev Biochem. 2002;71(1):473–510. 328. Backs J, Stein P, Backs T, et al. The gamma isoform of CaM kinase II controls mouse egg activation by regulating cell cycle resumption. Proc Natl Acad Sci U S A. 2010;107(1):81–86. 329. de Paola M, Bello OD, Michaut MA. Cortical granule exocytosis is mediated by alpha-­SNAP and N-­ethilmaleimide sensitive factor in mouse oocytes. PLoS One. 2015;10(8):e0135679. 330. Wang HH, Cui Q, Zhang T, et al. Rab3A, Rab27A, and Rab35 regulate different events during mouse oocyte meiotic maturation and activation. Histochem Cell Biol. 2016;145(6):647–657. 331. Mehlmann LM, Uliasz TF, Lowther KM. SNAP23 is required for constitutive and regulated exocytosis in mouse oocytes. Biol Reprod. 2019;101(2):338–346. 332. Tsai PS, van Haeften T, Gadella BM. Preparation of the cortical reaction: maturation-­ dependent migration of SNARE proteins, clathrin, and complexin to the porcine oocyte’s surface blocks membrane traffic until fertilization. Biol Reprod. 2011;84(2):327–335. 333. Familiari G. Structural changes of the zona pellucida during fertilization and embryo development. Front Biosci. 2008;13:6730–6751. 334. Que EL, Duncan FE, Bayer AR, et al. Zinc sparks induce physiochemical changes in the egg zona pellucida that prevent polyspermy. Integr Biol (Camb). 2017;9(2):135–144. 335. Garside WT, Loret de Mola JR, Bucci JA, Tureck RW, Heyner S. Sequential analysis of zona thickness during in vitro culture of human zygotes: correlation with embryo quality, age, and implantation. Mol Reprod Dev. 1997;47(1):99–104. 336. Drobnis EZ, Andrew JB, Katz DF. Biophysical properties of the zona pellucida measured by capillary suction: is zona hardening a . 1988;245(2):206–219.

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216.e8 PART I  The Fundamentals of Reproduction 337. Bleil JD, Beall CF, Wassarman PM. Mammalian sperm-­egg interaction: fertilization of mouse eggs triggers modification of the major zona pellucida glycoprotein. ZP2. Dev Biol. 1981;86(1):189–197. 338. Burkart AD, Xiong B, Baibakov B, Jiménez-­Movilla M, Dean J. Ovastacin, a cortical granule protease, cleaves ZP2 in the zona pellucida to prevent polyspermy. J Cell Biol. 2012;197(1):37–44. 339. Evans JP. Preventing polyspermy in mammalian eggs-­contributions of the membrane block and other mechanisms. Mol Reprod Dev. 2020;87(3):341–349. 340. Ravaux B, Favier S, Perez E, Gourier C. Egg CD9 protein tides correlated with sperm oscillations tune the gamete fusion ability in mammal. J Mol Cell Biol. 2018;10(6):494–502. 341. Moos J, Visconti PE, Moore GD, Schultz RM, Kopf GS. Potential role of mitogen-­activated protein kinase in pronuclear envelope assembly and disassembly following fertilization of mouse eggs. Biol Reprod. 1995;53(3):692–699. 342. Liu L, Yang X. Interplay of maturation-­ promoting factor and mitogen-­ activated protein kinase inactivation during metaphase-­ to-­interphase transition of activated bovine oocytes. Biol Reprod. 1999;61(1):1–7. 343. Scheffler K, Uraji J, Jentoft I, et al. Two mechanisms drive pronuclear migration in mouse zygotes. Nat Commun. 2021;12(1):841. 344. Fishman EL, Jo K, Nguyen QPH, et al. A novel atypical sperm centriole is functional during human fertilization. Nat Commun. 2018;9(1):2210. 345. Wong CC, Loewke KE, Bossert NL, et al. Non-­invasive imaging of human embryos before embryonic genome activation predicts development to the blastocyst stage. Nat Biotechnol. 2010;28(10):1115–1121. 346. Reichmann J, Nijmeijer B, Hossain MJ, et al. Dual-­spindle formation in zygotes keeps parental genomes apart in early mammalian embryos. Science. 2018;361(6398):189–193. 347. Mayer W, Smith A, Fundele R, Haaf T. Spatial separation of parental genomes in preimplantation mouse embryos. J Cell Biol. 2000;148(4):629–634. 348. Jefferson WN, Williams CJ. Early mouse embryo asymmetry. Mol Reprod Dev. 2012;79(7):433. 349. Sutovsky P, Moreno RD, Ramalho-­Santos J, Dominko T, Simerly C, Schatten G. Ubiquitin tag for sperm mitochondria: development. Nature. 1999;402(6760):371–372. 350. Rojansky R, Cha MY, Chan DC. Elimination of paternal mitochondria in mouse embryos occurs through autophagic degradation dependent on PARKIN and MUL1. Elife. 2016;5:e17896. 351. Song WH, Yi YJ, Sutovsky M, Meyers S, Sutovsky P. Autophagy and ubiquitin-­ proteasome system contribute to sperm mitophagy after mammalian fertilization. Proc Natl Acad Sci U S A. 2016;113(36):E5261–E5270. 352. Luo SM, Ge ZJ, Wang ZW, et al. Unique insights into maternal mitochondrial inheritance in mice. Proc Natl Acad Sci U S A. 2013;110(32):13038–13043. 353. Hutchison 3rd CA, Newbold JE, Potter SS, Edgell MH. Maternal inheritance of mammalian mitochondrial DNA. Nature. 1974;251(5475):536–538. 354. Vassena R, Boué S, González-­ Roca E, et al. Waves of early transcriptional activation and pluripotency program initiation during human preimplantation development. Development. 2011;138(17):3699–3709. 355. Xu YN, Shen XH, Lee SE, et al. Autophagy influences maternal mRNA degradation and apoptosis in porcine parthenotes developing in vitro. J Reprod Dev. 2012;58(5):576–584. 356. Toralova T, Kinterova V, Chmelikova E, Kanka J. The neglected part of early embryonic development: maternal protein degradation. Cell Mol Life Sci. 2020;77(16):3177–3194. 357. Tsukamoto S, Kuma A, Murakami M, Kishi C, Yamamoto A, Mizushima N. Autophagy is essential for preimplantation development of mouse embryos. Science. 2008;321(5885):117–120. 358. Yang Y, Zhou C, Wang Y, et al. The E3 ubiquitin ligase RNF114 and TAB1 degradation are required for maternal-­to-­zygotic transition. EMBO Rep. 2017;18(2):205–216. 359. Nishikimi A, Mukai J, Yamada M. Nuclear translocation of nuclear factor kappa B in early 1-­cell mouse embryos. Biol Reprod. 1999;60(6):1536–1541. 360. Knowland J, Graham C. RNA synthesis at the two-­cell stage of mouse development. J Embryol Exp Morphol. 1972;27(1):167–176. 361. Schramm RD, Bavister BD. Onset of nucleolar and extranucleolar transcription and expression of fibrillarin in macaque embryos developing in vitro. Reprod

362. Graf A, Krebs S, Heininen-­Brown M, Zakhartchenko V, Blum H, Wolf E. Genome activation in bovine embryos: review of the literature and new insights from RNA sequencing experiments. Anim Reprod Sci. 2014;149(1–2):46–58. 363. Dobson AT, Raja R, Abeyta MJ, et al. The unique transcriptome through day 3 of human preimplantation development. Hum Mol Genet. 2004;13(14):1461–1470. 364. Braude P, Bolton V, Moore S. Human gene expression first occurs between the four-­and eight-­cell stages of preimplantation development. Nature. 1988;332(6163):459–461. 365. Clegg KB, Pikó L. Quantitative aspects of RNA synthesis and polyadenylation in 1-­cell and 2-­cell mouse embryos. J Embryol Exp Morphol. 1983;74:169–182. 366. Bouniol C, Nguyen E, Debey P. Endogenous transcription occurs at the 1-­cell stage in the mouse embryo. Exp Cell Res. 1995;218(1):57–62. 367. Matsumoto K, Anzai M, Nakagata N, Takahashi A, Takahashi Y, Miyata K. Onset of paternal gene activation in early mouse embryos fertilized with transgenic mouse sperm. Mol Reprod Dev. 1994;39(2):136–140. 368. Park SJ, Komata M, Inoue F, et al. Inferring the choreography of parental genomes during fertilization from ultralarge-­scale whole-­ transcriptome analysis. Genes Dev. 2013;27(24):2736–2748. 369. Abe KI, Funaya S, Tsukioka D, et al. Minor zygotic gene activation is essential for mouse preimplantation development. Proc Natl Acad Sci U S A. 2018;115(29):E6780–E6788. 370. Zhu P, Guo H, Ren Y, et al. Single-­cell DNA methylome sequencing of human preimplantation embryos. Nat Genet. 2018;50(1):12–19. 371. Tahiliani M, Koh KP, Shen Y, et al. Conversion of 5-­methylcytosine to 5-­hydroxymethylcytosine in mammalian DNA by MLL partner TET1. Science. 2009;324(5929):930–935. 372. Gu TP, Guo F, Yang H, et al. The role of Tet3 DNA dioxygenase in epigenetic reprogramming by oocytes. Nature. 2011;477(7366):606–610. 373. Torres-­Padilla ME, Bannister AJ, Hurd PJ, Kouzarides T, Zernicka-­ Goetz M. Dynamic distribution of the replacement histone variant H3.3 in the mouse oocyte and preimplantation embryos. Int J Dev Biol. 2006;50(5):455–461. 374. van der Heijden GW, Dieker JW, Derijck AAHA, et al. Asymmetry in histone H3 variants and lysine methylation between paternal and maternal chromatin of the early mouse zygote. Mech Dev. 2005;122(9):1008–1022. 375. Ahmad K, Henikoff S. Histone H3 variants specify modes of chromatin assembly. Proc Natl Acad Sci U S A. 2002;99(suppl_4): 16477–16484. 376. Lin CJ, Conti M, Ramalho-­Santos M. Histone variant H3.3 maintains a decondensed chromatin state essential for mouse preimplantation development. Development. 2013;140(17):3624–3634. 377. Lin CJ, Koh FM, Wong P, Conti M, Ramalho-­Santos M. Hira-­ mediated H3.3 incorporation is required for DNA replication and ribosomal RNA transcription in the mouse zygote. Dev Cell. 2014;30(3):268–279. 378. Dahl JA, Jung I, Aanes H, et al. Broad histone H3K4me3 domains in mouse oocytes modulate maternal-­to-­zygotic transition. Nature. 2016;537(7621):548–552. 379. Zhang A, Xu B, Sun Y, et al. Dynamic changes of histone H3 trimethylated at positions K4 and K27 in human oocytes and preimplantation embryos. Fertil Steril. 2012;98(4):1009–1016. 380. Ooga M, Fulka H, Hashimoto S, Suzuki MG, Aoki F. Analysis of chromatin structure in mouse preimplantation embryos by fluorescent recovery after photobleaching. Epigenetics. 2016;11(1):85–94. 381. Ahmed K, Dehghani H, Rugg-­ Gunn P, Fussner E, Rossant J, Bazett-­Jones DP. Global chromatin architecture reflects pluripotency and lineage commitment in the early mouse embryo. PLoS One. 2010;5(5):e10531. 382. Abe KI, Yamamoto R, Franke V, et al. The first murine zygotic transcription is promiscuous and uncoupled from splicing and 3′ processing. EMBO J. 2015;34(11):1523–1537. 383. Zhang Y, Zhang X, Shi J, et al. Dnmt2 mediates intergenerational transmission of paternally acquired metabolic disorders through sperm small non-­coding RNAs. Nat Cell Biol. 2018;20(5):535–540. 384. Wu J, Xu J, Liu B, et al. Chromatin analysis in human early development reveals epigenetic transition during ZGA. Nature. 2018;557(7704):256–260. 385. Liu L, Leng L, Liu C, et al. An integrated chromatin accessibility implantation embryos.

CHAPTER 9  Meiosis, Fertilization, and Preimplantation Embryo Development 386. Li L, Guo F, Gao Y, et al. Single-­cell multi-­omics sequencing of human early embryos. Nat Cell Biol. 2018;20(7):847–858. 387. Ke Y, Xu Y, Chen X, et al. 3D chromatin structures of mature gametes and structural reprogramming during mammalian embryogenesis. Cell. 2017;170(2):367–381.e20. 388. Du Z, Zheng H, Huang B, et al. Allelic reprogramming of 3D chromatin architecture during early mammalian development. Nature. 2017;547(7662):232–235. 389. Chen X, Ke Y, Wu K, et al. Key role for CTCF in establishing chromatin structure in human embryos. Nature. 2019;576(7786):306–310. 390. LeGouy E, Thompson EM, Muchardt C, Renard JP. Differential preimplantation regulation of two mouse homologues of the yeast SWI2 protein. Dev Dyn. 1998;212(1):38–48. 391. Bultman SJ, Gebuhr TC, Pan H, Svoboda P, Schultz RM, Magnuson T. Maternal BRG1 regulates zygotic genome activation in the mouse. Genes Dev. 2006;20(13):1744–1754. 392. Jimenez R, Melo EO, Davydenko O, et al. Maternal SIN3A regulates reprogramming of gene expression during mouse preimplantation development. Biol Reprod. 2015;93(4):89. 393. Oldfield AJ, Henriques T, Kumar D, et al. NF-­Y controls fidelity of transcription initiation at gene promoters through maintenance of the nucleosome-­depleted region. Nat Commun. 2019;10(1):3072. 394. Lu F, Liu Y, Inoue A, Suzuki T, Zhao K, Zhang Y. Establishing chromatin regulatory landscape during mouse preimplantation development. Cell. 2016;165(6):1375–1388. 395. Hendrickson PG, Doráis JA, Grow EJ, et al. Conserved roles of mouse DUX and human DUX4 in activating cleavage-­stage genes and MERVL/HERVL retrotransposons. Nat Genet. 2017;49(6):925–934. 396. De Iaco A, Planet E, Coluccio A, Verp S, Duc J, Trono D. DUX-­ family transcription factors regulate zygotic genome activation in placental mammals. Nat Genet. 2017;49(6):941–945. 397. Leidenroth A, Clapp J, Mitchell LM, et al. Evolution of DUX gene macrosatellites in placental mammals. Chromosoma. 2012;121(5): 489–497. 398. Eckersley-­Maslin M, Alda-­Catalinas C, Blotenburg M, Kreibich E, Krueger C, Reik W. Dppa2 and Dppa4 directly regulate the Dux-­driven zygotic transcriptional program. Genes Dev. 2019;33(3–4):194–208. 399. De Iaco A, Coudray A, Duc J, Trono D. DPPA2 and DPPA4 are necessary to establish a 2C-­like state in mouse embryonic stem cells. EMBO Rep. 2019;20(5):e47382. 400. White MD, Bissiere S, Alvarez YD, Plachta N. Mouse embryo compaction. Curr Top Dev Biol. 2016;120:235–258. 401. Iwata K, Yumoto K, Sugishima M, et al. Analysis of compaction initiation in human embryos by using time-­lapse cinematography. J Assist Reprod Genet. 2014;31(4):421–426. 402. Ohsugi M, Hwang SY, Butz S, Knowles BB, Solter D, Kemler R. Expression and cell membrane localization of catenins during mouse preimplantation development. Dev Dyn. 1996;206(4):391–402. 403. Pauken CM, Capco DG. Regulation of cell adhesion during embryonic compaction of mammalian embryos: roles for PKC and beta-­ catenin. Mol Reprod Dev. 1999;54(2):135–144. 404. Fierro-­González JC, White MD, Silva JC, Plachta N. Cadherin-­ dependent filopodia control preimplantation embryo compaction. Nat Cell Biol. 2013;15(12):1424–1433. 405. Louvet S, Aghion J, Maria SA, Mangeat P, Maro B. Ezrin becomes restricted to outer cells following asymmetric division in the preimplantation mouse embryo. Dev Biol. 1996;177(2):568–579. 406. Dard N, Louvet-­Vallée S, Santa-­Maria A, Maro B. Phosphorylation of ezrin on threonine T567 plays a crucial role during compaction in the mouse early embryo. Dev Biol. 2004;271(1):87–97. 407. Maître JL, Niwayama R, Turlier H, Nédélec F, Hiiragi T. Pulsatile cell-­ autonomous contractility drives compaction in the mouse embryo. Nat Cell Biol. 2015;17(7):849–855. 408. Fleming TP. Cell adhesion in the preimplantation mammalian embryo and its role in trophectoderm differentiation and blastocyst morphogenesis. Front Biosci. 2001;6(1):d1000. 409. Sheth B, Fontaine JJ, Ponza E, et al. Differentiation of the epithelial apical junctional complex during mouse preimplantation development: a role for rab13 in the early maturation of the tight junction. Mech Dev. 2000;97(1–2):93–104. 410. Moriwaki K, Tsukita S, Furuse M. Tight junctions containing claudin 4 and 6 are essential for blastocyst formation in preimplantation mouse embryos. Dev Biol. 2007;312(2):509–522. 411. Biggers JD, Bell JE, Benos DJ. Mammalian blastocyst: transport functions in a developing epithelium. 1):C419–C432.

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412. Barcroft LC, Offenberg H, Thomsen P, Watson AJ. Aquaporin proteins in murine trophectoderm mediate transepithelial water movements during cavitation. Dev Biol. 2003;256(2):342–354. 413. Bell CE, Watson AJ. p38 MAPK regulates cavitation and tight junction function in the mouse blastocyst. PLoS One. 2013;8(4):e59528. 414. Manejwala FM, Schultz RM. Blastocoel expansion in the preimplantation mouse embryo: stimulation of sodium uptake by cAMP and possible involvement of cAMP-­dependent protein kinase. Dev Biol. 1989;136(2):560–563. 415. Hall JG. Twinning. Lancet. 2003;362(9385):735–743. 416. Knopman J, Krey LC, Lee J, Fino ME, Novetsky AP, Noyes N. Monozygotic twinning: an eight-­year experience at a large IVF center. Fertil Steril. 2010;94(2):502–510. 417. Kyono K. The precise timing of embryo splitting for monozygotic dichorionic diamniotic twins: when does embryo splitting for monozygotic dichorionic diamniotic twins occur? Evidence for splitting at the morula/blastocyst stage from studies of in vitro fertilization. Twin Res Hum Genet. 2013;16(4):827–832. 418. Dirican EK, Olgan S. On the origin of zygosity and chorionicity in twinning: evidence from human in vitro fertilization. J Assist Reprod Genet. 2021;38(11):2809–2816. 419. Sutherland K, Leitch J, Lyall H, Woodward BJ. Time-­lapse imaging of inner cell mass splitting with monochorionic triamniotic triplets after elective single embryo transfer: a case report. Reprod Biomed Online. 2019;38(4):491–496. 420. Derom C, Vlietinck R, Derom R, Van den Berghe H, Thiery M. Population-­based study of sex proportion in monoamniotic twins. N Engl J Med. 1988;319(2):119–120. 421. Sundaram V, Ribeiro S, Noel M. Multi-­chorionic pregnancies following single embryo transfer at the blastocyst stage: a case series and review of the literature. J Assist Reprod Genet. 2018;35(12):2109–2117. 422. Liu S, Hong Y, Cui K, et al. Four-­generation pedigree of monozygotic female twins reveals genetic factors in twinning process by whole-­ genome sequencing. Twin Res Hum Genet. 2018;21(5):361–368. 423. Sasaki H. Roles and regulations of hippo signaling during preimplantation mouse development. Dev Growth Differ. 2017;59(1):12–20. 424. Nishioka N, Inoue KI, Adachi K, et al. The hippo signaling pathway components lats and yap pattern tead4 activity to distinguish mouse trophectoderm from inner cell mass. Dev Cell. 2009;16(3):398–410. 425. Strumpf D, Mao CA, Yamanaka Y, et al. Cdx2 is required for correct cell fate specification and differentiation of trophectoderm in the mouse blastocyst. Development. 2005;132(9):2093–2102. 426. Niwa H, Toyooka Y, Shimosato D, et al. Interaction between Oct3/4 and Cdx2 determines trophectoderm differentiation. Cell. 2005;123(5):917–929. 427. Rayon T, Menchero S, Nieto A, et al. Notch and hippo converge on Cdx2 to specify the trophectoderm lineage in the mouse blastocyst. Dev Cell. 2014;30(4):410–422. 428. Galán A, Montaner D, Póo ME, et al. Functional genomics of 5-­to 8-­cell stage human embryos by blastomere single-­cell cDNA analysis. PLoS One. 2010;5(10):e13615. 429. Chen AE, Egli D, Niakan K, et al. Optimal timing of inner cell mass isolation increases the efficiency of human embryonic stem cell derivation and allows generation of sibling cell lines. Cell Stem Cell. 2009;4(2):103–106. 430. Nichols J, Zevnik B, Anastassiadis K, et al. Formation of pluripotent stem cells in the mammalian embryo depends on the POU transcription factor Oct4. Cell. 1998;95(3):379–391. 431. Chazaud C, Yamanaka Y, Pawson T, Rossant J. Early lineage segregation between epiblast and primitive endoderm in mouse blastocysts through the Grb2-­MAPK pathway. Dev Cell. 2006;10(5):615–624. 432. Kang M, Piliszek A, Artus J, Hadjantonakis AK. FGF4 is required for lineage restriction and salt-­and-­pepper distribution of primitive endoderm factors but not their initial expression in the mouse. Development. 2013;140(2):267–279. 433. Hashimoto M, Sasaki H. Epiblast formation by TEAD-­ YAP-­ dependent expression of pluripotency factors and competitive elimination of unspecified cells. Dev Cell. 2019;50(2):139–154.e5. 434. Huang TTF, Chinn K, Kosasa T, Ahn HJ, Kessel B. Morphokinetics of human blastocyst expansion in vitro. Reprod Biomed Online. 2016;33(6):659–667. 435. Fleming TP, Garrod DR, Elsmore AJ. Desmosome biogenesis in the mouse preimplantation embryo. Development. 1991;112(2):527–539. 436. Seshagiri PB, Sen Roy S, Sireesha G, Rao RP. Cellular and molecular regulation of mammalian blastocyst hatching. J Reprod Immunol.

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Structure, Function, and Evaluation of the Female Reproductive Tract Andrew M. Kelleher, Leah H. Bressler, Steven L. Young, and Thomas E. Spencer

OUTLINE STRUCTURE AND FUNCTION Ontogeny of the Uterus Role of the WNT Family and Homeobox Genes Steroid Action in the Endometrium Paracrine Actions of Steroid Hormones in the Endometrium Steroid Hormone Metabolism in the Endometrium MENSTRUAL CYCLE Histologic Changes ENDOMETRIUM IN THE CYCLE OF CONCEPTION AND PREGNANCY Secreted Proteins of the Endometrium Endometrial Preparation for Implantation Early Implantation Events Growth Factors and Cytokines Human Chorionic Gonadotropin Prostanoids and Other Lipids Immunology of the Endometrium ENDOMETRIAL MICROBIOME CLINICAL EVALUATION OF THE ENDOMETRIUM Endometrial Biopsy Endometrial Receptivity Biomarkers and Clinical Evaluation Global Gene Expression Patterns During the Window of Implantation Ultrasonography Sonohysterography Hysteroscopy ENDOMETRIUM IN ADVANCING AGE The fallopian tubes, endometrium, myometrium, and cervix function in concert to receive gametes, facilitate fertilization, support embryo growth, and ultimately orchestrate the timely expulsion of a mature fetus. The preparation of a reproductive tract conducive to pregnancy is governed by ovarian steroid hormones acting directly on their cognate receptors and indirectly through multiple steroid-­regulated growth factors, cytokines, and other extracellular and intracellular signaling molecules. This complex and redundant interaction between cells is made further complicated by bidirectional communications between the placenta trophoblast/trophectoderm and the endometrial cells. This chapter describes the structural and biochemical changes in the endometrium during the normal menstrual cycle and pregnancy, its clinical evaluation, and the pathophysiology of some relevant disorders related to endometrial function (see also Chapters 14, 25, and 26). The components of a receptive endometrium include the luminal epithelium, whose apical surface expresses cell adhesion molecules permitting adherence of the blastocyst; glandular epithelium, whose cells secrete substances that support the devel opment of the blastocyst and placenta; and decidualized stromal

cells and large granular lymphocytes that modulate trophoblast function and functional and structural modification of blood vessels. The interregulation of these components is orchestrated by the secretion and action of growth factors, growth factor-­binding proteins, angiogenic factors, cytokines, and an extracellular matrix that facilitates trophoblast invasion. The combinatorial actions of paracrine and endocrine factors, together with the extracellular matrix, promote trophoblast proliferation and development into the endometrium while controlling excessive invasion. Innate and adaptive immune functions, under the regulation of steroid hormones, collectively defend the reproductive tract environment against microbial invasion, but also must be modulated during pregnancy to allow the semiallogenic embryo and fetus to avoid rejection by the maternal host. The vascular system nourishes the endometrium in the initial receptive phase and is subsequently remodeled by invading trophoblasts to establish the placental blood supply. The coordinated contractile activity of the myometrium promotes sperm migration in a cycle of conception while also facilitating embryo transport prior to attachment. In the absence of conception, the functionalis portion of the endometrium is shed through a well-­controlled inflammatory-­ like reaction involving enzymes such as matrix metalloproteinases (MMPs), inflammatory cytokines, production of vasoactive substances, and uterine contractions, leaving behind the endometrial basalis, with its stem cells to allow regeneration. Specialized mechanisms ensure hemostasis and prevent scar formation. Through these mechanisms, a new and intact luminal endometrial surface is regenerated and prepared for the next round of oocyte release and potential fertilization.

STRUCTURE AND FUNCTION • The primary function of the endometrium is to provide a privileged site for blastocyst implantation and to provide an optimal environment for growth and development of the embryo/fetus and its associated placenta. • The cyclic differentiation of the endometrium depends on the actions of steroid hormones from the ovary including estrogen and progesterone. • The endometrium undergoes repetitive cycles of proliferation, differentiation, and menstruation, hundreds of times in a woman’s life, without apparent signs of aging. • Embryo implantation requires a complex series of endometrial changes to allow optimal receptivity to the embryo and to govern placental development. • Inflammatory signaling plays an important role in normal uterine function (menstruation and embryo implantation), but inappropriate inflammation in the endometrium may result in a phenomenon known as progesterone resistance, which appears to be involved in infertility and pregnancy loss.

Ontogeny of the Uterus The female reproductive tract is derived from the urogenital ridge, which, during week 6 of gestation, gives rise to paired mesodermal, paramesonephric tubes (the müllerian or paramesonephric

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ducts). Differentiation of the intermediate mesoderm precedes development of the gonads (ovaries/testicles), kidneys, urinary tract, and male/female reproductive tracts.1,2 Mesenchymal to epithelial transformation within the intermediate mesoderm gives rise to tubules that eventually form the male and female reproductive tracts (see Chapter 16).3,4,5 Longitudinal invaginations of the coelomic epithelium form the Fallopian tubes or oviducts, with the caudal ends fusing by week 10 of gestation to produce the primordial uterus and upper portion of the vagina. A thin septum remaining after fusion eventually resorbs, yielding a single uterine cavity. The primordial uterus is initially lined by a simple cuboidal epithelium that subsequently is specialized during postnatal development in domestic animals, laboratory rodents, and humans.6–9 Postnatal radial patterning establishes the major histological elements of the developed uterus, including stratification of the endometrial stroma, differentiation of the myometrium, and development and differentiation of the luminal (LE) and glandular epithelium (GE). Adenogenesis is initiated by differentiation of the GE from precursor LE. Nascent glands elongate into the stroma and then coil and slightly branch as they develop through the stroma toward the inner circular layer of the myometrium. By week 22 of gestation, the uterus has the primary structure of the adult organ. Glandular secretory activity, glycogen accumulation, and stromal edema are evident by week 32 under the influence of placentally derived steroid hormones. After delivery and the consequent precipitous fall in placental estrogens and progesterone, the endometrium regresses to an atrophic state, containing a few small glands and a poorly vascularized stroma.

Role of the WNT Family and Homeobox Genes The embryonic events described previously are driven, in large measure, by the secreted ligands of the wingless-­type MMTV integration site (WNT) family (WNT4, WNT5A, WNT7A) and transcriptional regulators of the homeobox (HOX) gene family based predominantly on studies in mice (Fig. 10.1).10,11 This morphogenetic program can only be played out in the absence of antimüllerian hormone (AMH also known as müllerian Inhibiting Substance or MIS) is a member of the transforming growth factor beta (TGFB) family made by the Sertoli cells of the fetal testes. In the absence of testosterone and AMH, the müllerian ducts elongate and develop into the fallopian tubes, uterus, cervix, and upper part of the vagina. The elongation phase of the müllerian ducts requires a number of factors. Given their common embryonic origin, early development in the mouse of the kidneys, ureters, and reproductive tract are tightly linked and involve other specific genes, including Pax2, Lim1, Emx2, as well as the members of the WNT and abdominal-­B HOXA families of genes.12 Lim1 encodes a transcription factor that along with PAX2 is essential for urogenital tract development.13 Lim1 null mice lack uteri and oviducts.14 Pax2 null mice lack kidneys, ureters, and genital tracts.15 Caudal elongation of the paramesonephric duct is absent. EMX2 is another transcription factor of the homeodomain gene family that appears to be essential for urogenital tract development.16,17 EMX2 is highly expressed in the adult uterus, and its expression is correlated with cell proliferation and appears to be inhibited by the HOX gene, HOXA10. There is decreased expression of PAX2 and LIM1, and mesenchymal segmental polarity gene product, WNT4, is also absent in mice lacking EMX2, suggesting the essential role of this transcription factor. The function of individual WNTs in uterine development has been addressed by targeted deletion of specific WNT genes. Wnt4 and Wnt5a are expressed throughout the mesenchyme of the FRT, whereas Wnt7a is expressed uniquely in LE, and crosstalk between these compartments is essential for uterine

development.10,18 The müllerian ducts are absent in female mice lacking Wnt4, a gene expressed in the mesenchyme.12 Moreover, female mice lacking WNT4 are partially sex-­ reversed due to the retention of the Wolffian ducts. Cases of Wnt4 null mutations associated with müllerian duct regression and a phenotype, including hyperandrogenemia, resembling that of the Wnt4 knockout mouse, have been reported in women.12 There is also a proposed role for WNT4 in postnatal uterine function, including progesterone signaling.19 WNT9B is expressed in the Wolffian duct epithelium and is necessary for müllerian duct extension.20 Mutations in the WNT9B gene have been found in women with Mayer-­Rokitansky-­Küster-­ Hauser (MRKH) syndrome.21 These müllerian defects have been reported to be associated with other gene defects, including PBX1, RBM8A, and TBX6 mutations.22–25 Other genes have been implicated as well, based on balanced translocation studies and breakpoint mapping.26 Deficiency of Wnt5a, a gene expressed in the genital tubercle and genital tract mesenchyme, results in mice with stunted genital tubercles and the absence of external genitalia.10 WNT7A expression gives rise to the luminal and glandular epithelium of the fallopian tubes and uterus,27 while WNT5 is associated with stromal cells.18,28 WNT7A is involved in paracrine signaling to the endometrial mesenchyme. Mesenchymal beta-­catenin (CTNNB1) appears to be the essential downstream effector of the WNT7A pathway and mediates its effects on the oviduct and proper development of the uterus.27 Although mutations in Wnt7a have not been found in women with müllerian anomalies,29,30 mice lacking Wnt7a develop an oviduct that is not clearly demarcated from the upper uterine horn, and the uterus develops cellular characteristics that are similar to the vagina (including a stratified epithelium without uterine glands), and the uterine smooth muscle is disorganized.10 Postnatal expression of HOXA10 and HOXA11 in the uterus is also lost. Mesenchymal CTNNB1 appears to be the essential downstream effector of the Wnt7a pathway and mediates its effects on the oviduct and the proper development of the uterus.31 The WNT family of genes, including receptors and downstream signaling molecules, are also expressed in a regulated fashion in the adult reproductive tract, indicating that they have additional roles beyond those involved in early morphogenetic events, including the regulation of steroid hormone action in adult tissues (discussed in subsequent chapter text).32,33 Formation and anterior-­ posterior patterning of the müllerian duct is regulated primarily by homeodomain-­containing transcription factors. The HOX genes encode an evolutionarily conserved family of transcription factors that contain a signature 60 amino acid DNA-­binding homeodomain.12 They play critical roles in organizing cells along the anterior-­posterior axis and directing them to select a particular pathway of development. Mammalian HOX genes are arranged in four different clusters, designated A through D, with each cluster organized in a linear arrangement that parallels the order of expression along the anterior-­posterior body axis. Expression of HOXA genes in the human and mouse reproductive tract is conserved, with HOXA9 being expressed in the fallopian tubes, HOXA10 and HOXA11 in the uterus, HOXA11 in the cervix, and HOXA13 in the upper vagina.12 Although there is a consistent regional distribution of HOX gene expression along the reproductive tract, there is evidence for some functional redundancy among the adjacent genes. Like the WNT genes, the HOXA genes are also expressed in the adult uterus, and their expression is under estrogen and progesterone regulation. The importance of the HOX gene family in reproductive tract function was demonstrated through targeted deletions in specific HOXA genes.34 Another significant discovery was that Hand–Foot–Genital syndrome and Guttmacher syndrome, autosomal dominant conditions that affect bones in the hands

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HOXA9

HOXA10

HOXA11

HOXA13

Tubes

Uterus

Cervix

Vagina

WNT4 WNT5A Mesenchyme

WNT7A Epithelial cell

and feet and cause reproductive tract abnormalities (including the bicornuate uterus), are caused by mutations in the HOXA13 gene.35,36 However, to date, mutations in HOXA7 to HOXA13 and HOX gene cofactor pre-­B-­cell leukemia homeobox1 (PBX1) have yet to be found in subjects with congenital absence of the uterus and vagina. HOXA10 and HOXA11 have both been implicated in the process of implantation.37,38 Mice with targeted deletions in the HOXA10 and HOXA11 genes have subtle abnormalities in uterine morphology, including transformation of the upper uterine segment into oviduct-­like histology (HOXA10 mutants), and reduced endometrial stromal development and expression of leukemia inhibitory factor (LIF) are seen in HOXA11 mutants. Notably, both HOXA10 and HOXA11 nullizygous females are infertile due to a uterine factor, implicating these genes in the implantation process in the adult. Mice lacking H6 homeobox 3 (Emx3), another HOX domain gene product, are also infertile due to an implantation defect associated with perturbations in WNT and LIF gene expression.16 Müllerian anomalies represent a complex collection of developmental defects occurring in up to 5% of the general population.39 Depending on the stage of development at which they occur, physical differences in the reproductive tract can be mild (e.g., a partial uterine septum) or severe, with complete absence of the cervix, uterus, and Fallopian tubes. These can be associated with infertility, endometriosis, and miscarriage. Some of the abnormalities require surgical correction and are often discovered at the time of puberty, if not before. Given the close developmental interaction between the müllerian and urinary system, it is not surprising that combined renal and müllerian duct anomalies occur.40 The recent clinical success of uterine transplantation provides a promising surgical treatment option for uterine factor infertility in extreme cases (see Chapter 40; Uterine Transplantation). Continued studies on the genetics of müllerian development will provide critical insights into the origins of reproductive tract anomalies.

Steroid Action in the Endometrium Many, but not all, of the uterine responses to steroid hormones are mediated by specific intracellular cognate receptors (see Chapter 5). These nuclear receptors serve as transcription factors undergoing striking spatial and temporal changes in expres sion during the menstrual cycle.

Fig. 10.1 Expression patterns of HOX and WNT genes in the female reproductive tract during development. (Modified from Taylor HS. The role of HOX genes on the development and function of the female reproductive tract. Semin Reprod Med. 2000;18:81–89.)

to steroid hormones is determined by the number of bioavailable hormones, which is influenced by hormone production rates as well as local steroid metabolism; the repertoire of steroid receptors, coactivators, and corepressors expressed; and the action of growth factors and cytokines that modulate the action of steroid hormone receptors.45–47 Postnatal patterning of the uterus is an ovarian steroid-­ independent event.48,49,50 The oviducts, uterus, cervix, and vagina form in mice with inactivating mutations of both nuclear estrogen receptors (ESR1 and ESR2).45 Despite this independence from maternal or fetal estrogens, normal differentiation of the female reproductive tract can, paradoxically, be disrupted by exogenous estrogens.46 Diethylstilbestrol (DES), a synthetic estrogen that causes uterine and cervical anomalies in exposed females (discussed later in this chapter), and polychlorinated biphenyls suppress expression of Wnt7a and alter the pattern of expression of HOXA9 and HOXA10 in the murine reproductive tract through ESR1.45–47 This suggests that alterations in HOXA and WNT gene expression are the likely molecular mechanism underlying the anatomical defects observed in human females exposed to DES in utero. Postnatal progesterone may also inhibit normal endometrial gland development, as shown in the neonatal ewe and mouse.51,52 Using this model, it appears that progesterone inhibition of gland development involves disruption of the WNT system,53 and illustrates that development may be independent of hormones but exposure at the wrong time may alter the normal developmental pathways.

Estrogen Receptor Signaling Estradiol is the primary trophic hormone for the uterus, mediating uterine growth through estrogen receptor alpha or ESR1, whose amounts are highest during the proliferative phase of the cycle and decline after ovulation in response to rising progesterone (Fig. 10.2).54 Immunohistochemical studies observed estrogen receptors in the nuclei of epithelial, stromal, and myometrial cells during the proliferative phase, with the epithelial cell staining being most prominent.42,43 After progesterone levels rise in the luteal phase, estrogen receptor staining is restricted to the deep basal glands and vascular smooth muscle. In situ hybridization studies demonstrate that mRNA transcripts for both ESR1 and ESR2 (coding for estrogen receptor alpha and beta, respectively) are expressed in the epithelial, stromal, and smooth muscle

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Fig. 10.2 Estrogen and progesterone receptors in endometrial cells during the normal menstrual cycle.  The concentrations of estrogen receptor (shaded area, A) and progesterone receptor (shaded area, B), mean values of plasma estradiol and progesterone (green lines, A and B), and endometrial proliferation as assessed by incorporation of [3H]thymidine in vitro (labeling index, C). The blue bar indicates menses. (Modified from Frolich M, Brand EC, van Hall EV. Serum levels of unconjugated aetiocholanolone, androstenedione, testosterone, dehydroepiandrosterone, aldosterone, progesterone, and oestrogens during the normal menstrual cycle. Acta Endocrinol [Copenh]. 1976;81:548–556.)

Is there a threshold dose of estrogen required to elicit a uterine growth response? Key and Pike55 proposed a threshold estrogen level of approximately 50 to 100 pg/mL, at which point endometrial proliferation is triggered and above which no further stimulation of endometrial proliferation occurs. This estimation is based on comparing endometrial proliferation assessed through ex vivo thymidine incorporation into endometrial explants from different stages of the menstrual cycle with corresponding estradiol levels at the different days of sampling.55 This hypothesis finds relevance in postmenopausal estrogen therapy where the effects of estrogen on bone, cardiovascular, and endometrial function may present differential risk/ benefit profiles.56

The decline in ESR1 at the time of implantation appears to be physiologically important57 and is a common finding across many species.43,44,58 Failure to decrease ESR1 indicates an imbalance in regulatory mechanisms of steroid hormone interactions and is associated with progesterone resistance, endometriosis, and infertility.59,60 The mechanism of this downregulation is complex, involving progesterone signaling that is discussed in more detail later in this chapter. ESR2 is expressed throughout the body expressed in almost all tissues.61 Despite a negligible effect on implantation in the knockout null mouse62, ESR2 does appear to have importance in endometrial function. Like ESR1, ESR2 does appear to be upregulated by estrogen and downregulated by progesterone.63 Furthermore, in the ESR2 knockout mouse, progesterone receptor (PGR) levels rose, suggesting a suppressive action of ESR2 on this receptor, and may also modulate ESR1.65 In Esr1 depleted mice, treatment with estrogen blocked proliferation, suggesting an antiestrogenic effect and causing epithelial apoptosis through ESR2.65 In the human endometrium, ESR2 has been reported to be expressed in both glands and stroma, but some suggest it is present exclusively in the vascular compartment.61 Relative overexpression of ESR2 in ectopic endometrium of women with endometriosis has also been reported. Paradoxically, in humans, this increase in ESR2 noted in endometriosis has been shown to stimulate the progression of disease.67 Other members of the estrogen receptor family include estrogen receptor-­related alpha (ESRRA) and beta (ESRRB), orphan receptors, with homology to the classical ESR1. ERRA and its coactivator peroxisome proliferator-­ activated receptor gamma (PPARG) coactivator-­1 alpha were found to show dramatically increased expression in the decidua and stimulate metabolic pathways for energy generation, perhaps in preparation for implantation.68 ERRB is observed throughout the endometrium during the menstrual cycle, including uterine natural killer (uNK) cells, but a precise role for this receptor has yet to be determined in the endometrium.69 Membrane-­ bound receptors with specific recognition of steroid receptors also coordinate the paracrine, autocrine, and juxtacrine cellular mechanisms and provide an explanation for the rapid effects of steroid hormones.51,53,70,71 Rapid effects of estradiol are mediated by at least two distinct receptors, a membrane-­associated form of ESR1 and a recently described integral membrane receptor, G protein-­ coupled estrogen receptor (GPER), previously known as G protein-­ coupled receptor 30 (GPR30).51,72 GPER has been characterized as an estrogen receptor and subsequent development of a GPER-­ specific agonist, G-­ 1, and antagonist, G-­ 15, have revealed important functions of this nonclassical estrogen receptor in multiple physiologic and pathophysiologic processes.73,74 In the endometrium, this receptor is present in the endometrial epithelium during the late proliferative phase and transitions to the stroma and decidua in the latter stages of the menstrual cycle and pregnancy but may be dysregulated in endometriosis.75,76 G-­1 has been shown to induce cell cycle arrest and thereby has some potential value in proliferative diseases such as endometriosis.77 GPER may also function as an aldosterone receptor, and though a mechanism conferring steroid specificity remains unclear.78

Progesterone Receptor Signaling Progesterone antagonizes the actions of estrogen in the endometrium and promotes differentiation of the glands and stroma via PGR. The antagonism of the uterotropic actions of estradiol involves a complex series of events, including alterations in estrogen receptor expression, inhibition of estrogen-induced translocation of the cell-cycle regulators, and induction of enzymes that catabolize estradiol.56,79,80 All

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of these effects are mediated through specific cognate nuclear receptors that are induced by estrogen and modulate downstream events in a paracrine fashion between the epithelial and stromal compartments. There are multiple isoforms of the PGR, but the majority of progesterone actions are served by the activities of either PGR-­ A or PGR-­ B. Both arise from a common PGR gene. Each isoform is expressed differentially in the endometrium, where the larger PGR-­B is a strong transcriptional activator of endometrial genes, while PGR-­A, 164 amino acid shorter, inhibits both PGR-­ B and other steroid receptors, including estrogen receptors. The A form predominates in the stroma, while B is more abundant in the epithelial phase of the early secretory phase of the cycle.81 Both PGR-­A and PGR-­B act as transcription factors, interacting with specific gene promoters and interacting with coregulators. PGR-­B also has nongenomic mechanisms of action, able to interact with Src tyrosine kinases in the cytoplasm, which independently modify gene expression.82–84 While generally considered to have opposing functions, PGR-­A is essential for stromal decidualization, and PGR-­B is downregulated in the epithelial compartment at the time of pregnancy. Indeed, the persistence of epithelial PGR is thought to be a sign of defective endometrial receptivity and may signify progesterone resistance.85 PGR isoforms A and B are both present in the epithelial and stromal compartments of the proliferative and early secretory

endometrium.86 Progesterone through PGR serves to inhibit proliferation by estrogen, primarily by downregulation of the estrogen receptor, but also through induction of estradiol degrading enzymes. Unlike its antiproliferative effects in the glandular epithelium, stromal PGR stimulates proliferation through activation of the MAPK/AKT pathway.87 Multiple other derived isoforms of PGR are thought to be present (Fig. 10.3).88–90 In addition to PGR-­A and PGR-­B, there is a highly truncated isoform PGR-­C, identified in the T47D breast cancer cell line.91 While still able to dimerize with PGR-­A or PGR-­B, PGR-­C may be an inhibitory factor, unable to interact with gene promoters. A role in labor for PGR-­C has been proposed.92 Classical endometrial PGR peak at the time of ovulation, localized to both epithelial and stromal cells, then declines. The rise in PGR is in response to estradiol, while the decline after ovulation is caused by elevated luteal phase progesterone that downregulates its own receptor.51,71,93 By 4 days after ovulation, PGR in the epithelial cells declines markedly in the epithelial compartment and remains weak or absent during the remainder of the secretory phase.43 In contrast, PGR expression in stromal cells remains strong throughout the menstrual cycle and into pregnancy, should it occur. In general, the A form predominates in the stroma, while the B is more abundant than the A form in the epithelial phase of the secretory phase of the cycle.81 PGR have not been detected in vascular endothelial cells or

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vascular smooth muscle, but they are abundant in the perivascular stroma. Consequently, the effects of progesterone or its withdrawal on the vasculature are likely indirect or perhaps via membrane PGR. The significance of different ratios of these two major PGR forms, with respect to regulation of gene expression in the human uterus, is increasingly being studied.94–96 Clues are being provided from the study of mice with targeted deletion of the receptor isoforms. The uteri of mice lacking both the A and B forms of the PGR are hyperplastic and contain inflammatory infiltrates.56,80 Hyperplasia reflects the lack of antagonism to the uterotropic actions of estradiol. Selective targeting of the A form of the PGR revealed that it is essential for progesterone-­mediated actions on implantation and the decidual response. However, the examination of genes associated with the window of uterine receptivity known to be regulated by progesterone indicated that the A form controls expression of only a subset, while others appear to be under the control of PGR-­B. Ablation of PGR-­A in mice uncovered an unexpected role for the B form of the receptor in inducing epithelial proliferation. Administration of estradiol to PGR-­A receptor-­deficient mice resulted in uterine hyperplasia, but the combination of estradiol and progesterone resulted in even greater hyperplasia. It thus appears that PGR-­A antagonizes the uterotropic responses mediated by both ESR1 and PGR-­B. Regarding implantation and decidualization, however, when PGR-­A null mice are examined, they are similar in their defects to mice that lack both PGR-­A and PGR-­B subunits, while PGR-­B knockout mice were fertile.97 Clinically, reductions in PGR-­B have been associated with proliferative states such as endometriosis,98 supporting this paradigm of counterregulatory mechanisms involving PGR isoforms. In addition, like estrogen, nongenomic actions of progesterone appear important in the reproductive tract as well. Membrane forms of PGR have also been identified in the uterus.84,84A This family of nonclassical, membrane progesterone receptors is structurally unrelated to PGR-­A, B, or C, but each of the mammalian paralogues has been shown to specifically bind progesterone and can activate G-­protein coupled signaling pathways.99 Indeed, membrane PGR (PGRMC) has been shown to rapidly activate MAP kinase and inhibit cyclic adenosine monophosphate (cAMP).101 Membrane PGR have been localized to the myometrium of humans and function to inhibit cAMP with a possible role of facilitating uterine contractions at term.101 In addition to steroid hormone receptors, coactivators and chaperone proteins have an important impact on the uterine response to estrogens and progestogens.45 Uterine expression of p160 coactivators, steroid receptor coactivator-­ 1 (SRC1), steroid receptor coactivator-­ 2 (SRC2), and steroid receptor coactivator-­ 3 (SRC3) has been examined.47 Decidualization of the endometrium does not occur without SRC2. SRC3 levels increased in the glandular epithelium in the late secretory phase, whereas SRC1 and SRC2 expression did not change. SRC3 has been linked to endometrial hyperplasia and in women with polycystic ovary syndrome (PCOS), SRC2 and 3, along with Erα, were elevated in the stroma and glandular epithelium,45 demonstrating that an abnormal endocrine milieu can alter coactivator levels, which could, in turn, result in endometrial dysfunction.

Paracrine Actions of Steroid Hormones in the Endometrium Based on the pioneering work of Cunha in mice,94,95 the effects of estrogen and progesterone on epithelial and stromal proliferation and differentiation are, in large part, indirect, involving paracrine substances produced by the stroma that act on the epithelium.96,102,

promotes DNA synthesis in the epithelium; under the influence of progesterone, the epithelium produces substances that affect the response of the underlying stroma and the epithelial cells to estrogen. These indirect actions of estradiol on epithelial proliferation have been demonstrated in elegant reconstitution and grafting experiments employing stroma and epithelium from normal and Esr1 knockout (ERKO) mice. Epithelial cell proliferation does not occur when the stroma from an ERKO mouse uterus is paired with epithelium from a normal uterus.96 Conversely, epithelial cell DNA synthesis in response to estrogen occurs when normal stroma is paired with epithelium from ERKO mice. Studies on human endometrial cells in culture are consistent with the mouse studies.104 Estradiol increases epithelial cell proliferation when cocultured with stroma, but it does not increase proliferation in epithelial cells cultured in the absence of stromal cells. What is the estrogen-­ stimulated signal from the stroma that promotes epithelial cell proliferation? Candidates include growth factors that are transcriptionally regulated by ESR1, including insulin-­ like growth factor-­ 1 (IGF1), transforming growth factor alpha (TGFA), and epidermal growth factor (EGF).105 Alternatively, estradiol might suppress production of stromal factors that restrain epithelial cell proliferation. Among the growth factors, there has been particular interest in EGF and IGF1. Studies using mouse models, including transplantation of uteri and vagina from EGF receptor knockout mice, indicate that this receptor is required for the maximal fibromuscular stroma growth but not the epithelial cell proliferative response to estrogen.106 IGF1 and IGF2 both stimulate the proliferation of human endometrial stromal cells via the type 1 IGF receptor. IGF1 expression is greatest in the late proliferative and early secretory phase, whereas IGF2 is most abundant in the midsecretory endometrium and decidua of the first trimester of pregnancy. The IGFs are bound by a family of binding proteins (IGFBPs) that modulate IGF activities in target tissues. One of the binding proteins, IGFBP1, is a major secretory product of decidualized stromal cells, and it has been hypothesized to play a role in controlling trophoblast invasion.107 Estrogen is the primary regulator of IGF1 expression in the uterus, which occurs predominantly in the stroma.108,109 Estrogen also increases expression of IGF1 receptors, which are primarily found on epithelial cells. The mitogenic response of the mouse uterus to estrogen is absent in mice with targeted deletion of the IGF1 gene. Moreover, mice overexpressing IGFBP1, which results in reduced IGF1 bioavailability, have a blunted DNA synthesis response to estrogen treatment.110 Thus, IGF1 produced in the uterine stroma in response to estrogen acts on the epithelial cells to stimulate DNA synthesis; however, tissue grafting experiments showed that an IGF1 knockout mouse uterus responds to estrogen when placed into a normal mouse, whereas a wild-­type uterus placed into an IGF1 knockout mouse showed minimal growth—indicating that systemic IGF1 is sufficient to support estrogen-­driven uterine growth.111 These findings substantiate the importance of IGF1 in the uterine growth response to estrogen. Although uterine growth can occur in the absence of a paracrine IGF-­1 system, these studies do not preclude a role for locally generated IGF1 as a redundant signaling mechanism or the role of other locally produced growth factors. Progesterone is a vital steroid hormone that is involved in secretory preparation of the endometrium for implantation, decidualization, and suppression of myometrial contractility during pregnancy.112 Similar to estrogen, progesterone effects on the endometrium involve paracrine activities.113,114 Progesterone possesses antiinflammatory characteristics and promotes immunotolerance during implantation and pregnancy.115 It is also associated directly, or indirectly, with most of the secretory proteins made by the endometrium that are

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present in the uterine lumen and glands to support embryo development and implantation.116 Progesterone signaling in the endometrium affects each of the cellular compartments, including the epithelium, stroma, and resident immune cells.117,118 It has become increasingly clear that progesterone has both direct and important indirect roles in both stroma and epithelial function.116 A primary role of progesterone after ovulation is to downregulate the actions of estrogen, which are unnecessary for normal secretory endometrial development119,120 and may, in fact, inhibit implantation.57 During the early proliferative phase, when progesterone concentrations are low, cytoplasmic PGR is shielded from degradation by heat shock proteins and by interactions with coactivator proteins (SRC1 and SRC2) and chaperone proteins such as FKBP4 and FKBP5. In SRC2 conditional knockout mice), the females are infertile, similar to FKBP4 null mice, with each displaying implantation and decidualization failure.117,118 In women, both FKBP4 and FKPB5 expression normally increases in the secretory phase, possibly regulated by HOXA10. Interestingly, blunted increases in FKBP4 are seen in endometrium from women with infertility and endometriosis,121 apparently regulated in part by changes in microRNA that regulate the degradation of mRNA coding for FKBP4 and other progesterone-­regulated genes.122–124 This mechanism may be involved in the phenomenon of endometrial progesterone resistance.125 Much of our understanding of implantation and the paracrine actions of progesterone is based on studies in the mouse uterus. PGR is upregulated by estrogen but also requires the transcription factor GATA2.126 The action of progesterone begins with epithelial PGR binding to progesterone followed by the induction of a key epithelial target gene, Indian hedgehog (IHH).114 IHH is secreted and communicates with the endometrial stroma in a paracrine fashion, binding to the stromal receptor Patched-­1 (Ptch1), resulting in increased stromal transcription factor, chicken ovalbumin upstream promoter transcription factor II (COUP-­TFII or NR2F1), with downstream effectors including bone morphogenic protein-­2 (BMP2) and WNT4. This pathway concludes with the induction of HAND2 in the stroma that is antiproliferative by inhibiting production of fibroblast growth factors (FGFs), which are required for the expression of epithelial estrogen receptors (Fig. 10.4).127 As mentioned earlier in this chapter, WNT4 is required for müllerian development and in adults is required for decidualization. WNT4 is also thought to be responsible for shuttling Forkhead box protein 1 (FOXO1), a transcription factor essential for decidualization,119 from the nucleus to the cytoplasm, preventing its apoptotic actions. Inadequate decidualization has been shown to be associated with implantation failure and associated with many critical genes.128 Many of the stromal genes expressed in response to progesterone action include HOXA10, heparin-­binding EGF-­like growth factor (Hbegf), cyclooxygenase 2 (Cox-­2, encoded by Ptgs2), and mitogen inducible gene 6 (Mig-­6 or Errfi1), are induced through the IHH pathway. Mig-­6 is a negative regulator of EGF receptor, and transgenic knockout mice for this protein have endometrial hyperplasia, suggesting this progesterone-­regulated gene is an important brake for cell proliferation in the endometrium. The HOX gene, HOXA10, is essential for uterine development, but in the adult has been shown to regulate FKPB4129 and is therefore also essential for progesterone action and implantation. HOXA10 also directly upregulates epithelial receptivity genes such as the ανβ3 integrin,130 which was shown to be required for attachment of the embryo in animal models.131,132 Mice lacking PGR, HOXA10, PTGS2, ARID1A, LIF, and other genes in transgenic mice have been found to exhibit decidualization defects and infertility.130,133,134 The uterus is also a target for androgens.135 Androgen recep tors (AR) are expressed in the endometrium and myometrium,

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Fig. 10.4 Progesterone signaling in the endometrium involves communication between the epithelium and the stroma, with complex interactions of the Indian Hedgehog and WNT pathways. One of the down-­stream effects of progesterone is the down-­regulation of estrogen signaling in both endometrial epithelial and stromal cells. (Modified from Large MJ, Demayo FJ. The regulation of embryo implantation and endometrial decidualization by progesterone receptor signaling. Mol Cell Endocrinol. 2012;358:155–165.)

most prominently in the stromal cells during the proliferative phase and in the epithelial cells of the secretory phase. MAGE-­ 11 was originally described as a primate-­specific AR coactivator,136 reaches peak expression in the secretory phase, and colocalizes with AR.88 Interestingly, 5 alpha-­reductase types 1 and 2, which convert testosterone to dihydrotestosterone, are expressed in epithelial cells throughout the cycle.136 Estradiol treatment increases endometrial stromal AR expression in the Rhesus monkey, and estradiol in combination with either testosterone or progesterone augments epithelial and myometrial AR levels. Consistent with these observations, AR expression is elevated in the endometrium of women with PCOS; this finding may explain, in part, the implantation defects and early pregnancy loss reported to be associated with this syndrome.137 Androgens and AR may also have essential roles in reproductive tract function in females, and this topic is becoming an active area of research.138

Steroid Hormone Metabolism in the Endometrium The activity of steroid hormone in the endometrium is determined in part by the modulating effects of uterine enzymes that actively catabolize steroid hormones (Fig. 10.5; see Chapter 4).139 These enzymes that carry out transformations of steroid hormones are subject to regulation during the menstrual cycle. Estradiol taken up from the plasma can be converted into estrone by the action of 17-­beta hydroxysteroid dehydrogenases (HSD17B1) or converted to sulfated conjugates by estrogen sulfotransferase.140 Three different forms of HSD17B capable of oxidizing estradiol to estrone have been detected in primate endometrium: the type 2, type 4, and type 8 enzymes.141 Type 2 and type 8 enzymes are associated with microsomes; type 4 enzyme is in peroxisomes. Type 2 and type 4 enzymes use the oxidized form of nicotinamide adenine dinucleotide as a cofactor and are predominantly localized to the glandular epithelium in the secretory phase. The endometrial type 2 enzyme shows the greatest change in expression

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during the cycle, peaking in the secretory phase. The type 8 enzyme appears to be constitutively expressed. Progesterone enhances the conversion of estradiol to estrone in endometrial cells by inducing expression of the type 2 HSD17B and, to a much lesser degree, the type 4 enzyme. Progesterone also increases endometrial estradiol sulfation by increasing estrogen sulfotransferase activity. Thus progesterone activates enzymatic pathways that inactivate estradiol. However, there are regional differences in the balance of systems removing and restoring estradiol in the uterus. For example, estrogen sulfatase is detected only in the glandular epithelium of the basalis, where it may function to increase the level of estradiol from estradiol sulfate.142 The human uterus does not normally have the capacity to produce significant amounts of estrogen locally (either de novo or from circulating prohormones), but endometrial cancers, endometriotic lesions, and to a lesser extent, eutopic endometrium of women with endometriosis aberrantly express components of the steroidogenic machinery (P450arom) that endows the tissue with the capacity to synthesize estrogens from circulating adrenal or ovarian androgens or by de novo synthesis. Endometriosis lesions have been found to express StAR, the cholesterol side-­chain cleavage enzyme, P450c17, aromatase, and type 1 HSD17B in stromal cells (see Chapter 25). Estradiol produced in these lesions can enhance the production of prostaglandin E2, which in turn stimulates transcription of the aromatase gene, resulting in a feedforward mechanism for increasing local estrogen levels. In addition, endometriotic lesions do not express the progesterone-­regulated type 2 HSD17B that converts estradiol to estrone, which effectively increases the bioavailability of estradiol. Progesterone is catabolized in the uterus into inactive 20α-­ hydroxyprogesterone by 20α-­HSDs. The type 2 17β-­HSD, which is increased in the secretory phase, is also a 20α-­hydroxysteroid oxidase that converts 20α-­hydroxyprogesterone back into progesterone.139 The induction of the type 2 HSD17B by progesterone in the secretory phase, therefore, contributes not only to the catabolism of estradiol but also to the maintenance of endome trial progesterone levels.

Fig. 10.5 Steroid metabolism and action in the endometrium. P4 prevents the E2-­stimulated translocation of cyclin Di and Cdk4 into the nucleus while activating Cdk2, a cyclin A kinase, which inhibits the mitogen effects of E2. +, Stimulation; −, inhibition; E1, estrone; E1S, estrone sulfate; E2, estradiol; E2R, estrogen receptor; E2S, estradiol sulfate; HSD, hydroxysteroid dehydrogenase; P4, progesterone; P4R, progesterone receptor; T, testosterone.

MENSTRUAL CYCLE The endometrium undergoes cyclic changes in response to preovulatory and postovulatory steroid hormones, to prepare for blastocyst implantation and pregnancy. In discussions of structure and function, the primate endometrium is commonly described as consisting of two major layers, the functionalis, and basalis (Fig. 10.6). The functionalis is a transient layer consisting of a compact zone that includes the stroma subjacent to the luminal epithelium and an intermediate spongy zone containing more densely packed glands. The basalis, or basal layer, lies beneath the spongy zone and adjacent to the myometrium. It contains the gland fundi and supporting vasculature and can regenerate the functionalis endometrium after it is shed at menstruation. These endometrial layers are histologically definable during the secretory phase. Through the menstrual cycle, the functionalis undergoes striking transformations, whereas the basalis remains relatively unchanged. Patterns of cell proliferation, programmed cell death, and gene expression also show gradients across the layers (described below). The majority of epithelial cell proliferation occurs in the upper regions of the functionalis, whereas proliferative activity in glands of the basalis is modest during the proliferative phase. Increased proliferative activity is observed in the midsecretory phase in the basalis layer.143–145 Of note, the epithelial cells of the basalis later maintain both ESR1 and PGR at a time when these receptors are normally depleted in the upper functionalis epithelium during the midsecretory phase.43

Histologic Changes Early Proliferative Phase During the early proliferative phase, the endometrium is usually less than 2 mm in thickness. Proliferation of cells in the basal zones and epithelial cells persisting in the lower uterine and cornual segments results in the restoration of the luminal epithelium by day 5 of the menstrual cycle. At that time, mitotic activity is evident in both the glandular epithelium and stroma. Remarkably,

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Fig. 10.6 Histologic organization of secretory-­phase human endometrium. (Modified from Weiss L, Creep RO, eds. Histology. 4th ed. McGraw-­Hill; 1977:911.)

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this recurrent “wound healing” process does not normally produce scarring. Endometrial stem cells capable of yielding progenitors of both the stromal and epithelial components of the endometrium presumably contribute to the regenerative process but are yet to be convincingly identified.146 Rapid rejuvenation of the endometrium depends on many of the same factors that were involved in the ontogeny of the reproductive tract. WNT7A is expressed solely by the luminal epithelium and is a diffusible secreted factor that triggers cell proliferation through complex pathways.147,148 Acting on the underlying stroma, soluble WNT7A binds to the receptor Frizzled (FZL) that phosphorylates the intracytoplasmic protein Disheveled (DSH; Fig. 10.7A). This protein inactivates glycogen synthase kinase beta (GSKB), turning off the breakdown of CTNNB1 by ubiquitination. Accumulation of CTNNB1 signals cell proliferation activities associated with endometrial growth acting as a transcription factor in the nucleus. The diffusion gradient of WNT7A downward into the growing endometrium is an attractive model for self-­regulatory mechanisms to determine the growth of the endometrium. Counterregulatory mechanisms to disable WNT7A/FZL/ DSH pathways include the action of progesterone, which stimulates the secretion of a protein called Dickkopf-­1 (DKK1; see Fig. 10.7B).147,148 DKK1 binds to a coreceptor, LRP6, blocking the FZL receptor, turning off the action of WNT7A by preventing CTNNB1 accumulation. Defects in DKK1 production have been described in endometriosis and reflect progesterone resistance and might explain the proliferative phenotype of the endometrium in this condition.121,149 The glands during the early proliferative phase are narrow, straight, and tubular, lined with low columnar cells that have round nuclei located near the cell base (Fig. 10.8). At the ultrastructural level, the epithelial cell cytoplasm contains numerous polyribosomes, but the endoplasmic reticulum and Golgi com plexes of these cells are not well developed.

Late Proliferative Phase The endometrium thickens in the late proliferative phase due to glandular hyperplasia and an increase in the stromal extracellular matrix. The glands are widely separated near the endometrial surface and more crowded and tortuous deeper into the endometrium. The glandular epithelial cells increase in height and become pseudostratified as the time of ovulation approaches (see Fig. 10.8D). The effect of steroid hormones on the proliferation and secretion within the endometrium is highly dependent on the zones (basalis vs. functionalis layers). Studies from the rhesus macaque endometrium using specific labeling techniques show proliferation during the “proliferative” phase is confined to the functionalis layer.150

Early Secretory Phase Ovulation marks the beginning of the secretory phase of the endometrial cycle. However, it should be noted that the endometrial luminal and glandular epithelial cells also display secretory activity during the proliferative phase. Mitotic activity in epithelial and stromal cells is restricted to the first 3 days after ovulation and is rarely observed later in the cycle. The nuclei of glandular epithelial cells and stromal cells develop heterochromatin in the early secretory phase (see Fig. 10.8). The glandular epithelial cells begin to accumulate glycogen-­rich vacuoles at their base, displacing the nuclei to the midregions of the columnar cells. Ultrastructural studies of endometrial epithelia reveal abundant endoplasmic reticulum and unusually large mitochondria with prominent cristae. A reticular network of argyrophilic fibers containing fibrillar collagens (collagen fiber type I and type III) is established in the stroma during this phase. Stromal edema contributes to the thicken-

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WNT7A

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B Fig. 10.7 WNT signaling pathways in proliferative and secretory phase of the menstrual cycle. WNT7A is a soluble factor thought to arise from the luminal epithelium. (A) Under conditions of estrogen action, WNT7A binds to the receptor, frizzled (FZL), initiating signaling through the intracytoplasmic protein disheveled (DSH). Disheveled binds axin and glycogen synthase kinase β (GSK-­β), allowing β-­catenin (β-­CAT) to accumulate and stimulate gene transcription leading to cell proliferation. (B) When progesterone levels increase, cellular dickkopf-­1 accumulate and inactivate Fzl through an accessory receptor, LRP. Without WNT7A, GSK facilitates the ubiquitination of β-­CAT, which makes it unavailable as a transcription factor.

Midsecretory Phase A characteristic feature of this phase of the cycle is the development of the spiral arteries. These vessels become increasingly coiled as they lengthen more rapidly than the endometrium thickens. The endometrial glands are tortuous in the midsecretory and late secretory phases. Their secretory activity reaches a maximum about 6 days after ovulation, as reflected by the loss of vacuoles from the epithelial cell cytoplasm (see Fig. 10.8E and F). The nucleolar channel system, an ordered spherical stack of interdigitating tubules, appears transiently in the nucleoli of approximately 5% to 10% of the secretory phase epithelial cells between days 16 and 24.151 The nuclear channel system is thought to form from an invagination of the inner nuclear membrane, providing a direct connection to the perinuclear space for the transport of mRNA to the cytoplasm. Nucleolar and coiled-­body phosphoprotein 1 (Nolc1 or Nopp140), a highly phosphorylated protein that associates with small nucleolar ribonucleoprotein particles required for RNA processing, appears to induce the formation of this intranuclear endoplasmic reticulum.152 The nucleolar channel system forms in response to progesterone and is an ultrastructural hallmark of the secretory phase during the expected time of implantation. Stromal cells around blood vessels enlarge and acquire an eosinophilic cytoplasm and a pericellular extracellular matrix in the mid-­to late secretory phase. These changes, referred to as

predecidualization to distinguish them from the further transformation of the stroma that occurs in a fertile cycle, subsequently spreads, accentuating the demarcation between the subepithelial compact zone and the spongy zone. Unlike many laboratory animal species, an embryonic signal is not required for initiation of decidualization in the human uterus. The fact that the predecidual changes occur first near blood vessels suggests that humoral or local factors provoke them. Among the local factors may be interactions with decidual granular lymphocytes, also referred to as uterine natural killer (uNK) cells. uNK cells encircle arterioles and closely associate with stromal cells through contacts that are remarkably similar to gap junctions.153 At the ultrastructural level, the predecidual stromal cells display well-­developed Golgi complexes and parallel lamellae of the endoplasmic reticulum. Their surrounding matrix consists of laminin, fibronectin, heparan sulfate, and type IV collagen.139,140 Substantial changes in gene expression also occur in stromal cells during decidualization.141 The stromal cells of the midsecretory and late secretory phase also express a repertoire of proteins that promote hemostasis, including tissue factor (TF), a membrane-­ associated protein that initiates coagulation when it contacts blood, and plasminogen activator inhibitor type 1 (PAI1), also known as Serpin E1, which restrains fibrinolysis.142,154,155 PAI1 may prevent focal hemorrhage that might result from trophoblast invasion during

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Premenstrual and Menstrual Phase The main histologic features of the premenstrual phase are degradation of the stromal reticular network, which is catalyzed by MMPs; infiltration of the stroma by polymorphonuclear and mononuclear leukocytes; and “secretory exhaustion” of the endometrial glands, whose epithelial cells now display basal nuclei. Morphologic changes in the nuclei of granular lymphocytes, including pyknosis and karyorrhexis suggesting apoptosis, have been proposed to be some of the first events presaging menses; these changes occur prior to the breakdown of extracellular matrix and leukocyte infiltration.156 In the glandular epithelium, the nucleolar channel system and giant mitochondria characteristic of the early and midsecretory phases have vanished. The endometrium shrinks preceding menstruation, partly because of diminished secretory activity and the catabolism of the extracellular matrix. A broader discussion of menstruation in the context of abnormal uterine bleeding (AUB) is discussed later in this chapter.

Menstruation Menstruation, caused primarily by progesterone withdrawal, marks a failure to achieve pregnancy and the need to shed the specialized uterine lining that results from spontaneous

Fig. 10.8 Histology of the endometrium during the menstrual cycle. (A and B) Proliferative endometrium. Mitoses are present (arrow). Nuclei in the glandular epithelium are pseudostratified. (C and D) Secretory endometrium, day 18. Subnuclear glycogen vacuoles are uniformly present in glandular epithelium (arrowheads). (E and F) Midsecretory endometrium. Glandular secretory activity is present, and the stroma is edematous. The stromal cells in the more superficial layers as well as around vessels have become pseudodecidualized and exhibit a flattened, polygonal configuration with distinct cell borders.

decidualization.157 The uniqueness of this process is highlighted by the fact that, although circulating progesterone and estrogen levels decline with corpus luteum regression in nonfertile cycles in all mammals, menstruation appears almost exclusively in humans and some Old World primates. In menstruating species, moreover, tissues that respond to estrogen and progesterone such as the fallopian tubes, vagina, and breast do not shed as ovarian steroid levels decline. The molecular mechanisms triggered by progesterone withdrawal include activation of the nuclear factor kappa beta (NFKB) transcriptional pathway (a major target of cytokines) and the resulting expression of genes like endometrial bleeding-­associated factor (EBAF), an anti-­TGFB cytokine that interferes with the action of other members of the TGFB family that promote endometrial integrity. This orchestrated blockade of the actions of TGFB appears to initiate many of the subsequent events of menstruation, including the elaboration of MMPs.158 There have been two major models describing the mechanisms driving menstruation. One of these is vasoconstriction-­ induced hypoxia and reperfusion. The functional zone of the human endometrium is supplied by spiral arterioles that, in contradistinction to the radial and basal arteries that feed them, are highly sensitive to steroid hormones. The classic studies of Markee159 utilized autologous endometrial transplants into the anterior eye chamber of the rhesus macaque, allowing direct examination of the endometrium through the cycle. These

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studies demonstrated that vasoconstriction of the arterioles and spiral arteries precedes the onset of menstrual bleeding by 4 to 24 hours. It has been proposed that bleeding occurs after the arterioles and arteries relax, leading to hypoxia or reperfusion injury. However, later endometrial perfusion studies failed to reveal any significant reduction in endometrial blood flow in the perimenstrual phase or during menstruation.160 In addition, an analysis of expression and localization of hypoxia-­inducible factor (HIF), a heterodimeric transcription factor induced by hypoxia and thus a biochemical marker of reduced oxygen availability, was unrevealing. No upregulation of the two HIF component subunits, HIF1A and HIF1B, was observed, and no nuclear localization of HIF subunits took place in the perimenstrual human endometrium.161 While these studies do not exclude the possibility of localized regions of vasoconstriction and hypoxia, they do bring into question the hypoxia/reperfusion model. An alternative hypothesis to the vasoconstriction model is that menstruation is a controlled inflammatory response engendered by the withdrawal of progesterone. The inflammation hypothesis is supported by two features: the prominent accumulation of leukocytes in the endometrium in the premenstrual phase153; and the release of matrix-­degrading enzymes characteristic of an inflammatory response.162–165 The hypothesis of progesterone withdrawal-­induced inflammation is supported by the uterine inflammation observed in mice lacking PGR. Apoptotic cell death, which can be triggered by inflammatory mediators, occurs in the late secretory phase first in stromal cells and then gradually spreads throughout the functionalis.166–168 Rescue from apoptosis has been shown to occur in vivo with the administration of progesterone or exogenous human chorionic gonadotropin (hCG).169 Changes in proteins involved in apoptosis appear to contribute to the regional programmed death in the endometrium. The antiapoptotic protein, BCL2, is prominently expressed in the glandular epithelium during the proliferative phase; expression declines in the secretory phase to reach low levels in the late secretory phase when apoptosis occurs.168 Studies report an inverse pattern of expression of survivin, a recently discovered inhibitor of apoptosis. Survivin binds to and blocks the effector cell death proteases, caspase-­3 and -­7. The activities of caspases-­3, -­8, and -­9 are higher in the secretory phase. Low rates of survivin expression in the glandular epithelium are found in the proliferative phase, rising to peak expression in the late secretory phase. The protein is localized to the nuclei of cells in the functionalis and the cytoplasm in cells in the basalis. This differential distribution may indicate that survivin is not capable of suppressing apoptotic death in the late secretory phase functionalis but performs this role in the basalis, consistent with the observed patterns of apoptotic cell death. Elevated levels of survivin in endometriotic lesions correlate with reduced apoptotic death of cells in these lesions.170 Although the vasoconstriction and inflammation hypotheses of menstruation might appear to be distinct, there are several overlapping biochemical features of hypoxia and inflammation, including the release of proinflammatory cytokines and apoptotic cell death that tend to blur the distinctions between these models. The vascular changes in the endometrium in the perimenstrual phase, resulting either from ischemia/hypoxia or from an inflammatory reaction, lead to extravasation of blood. Autophagy and heterophagy are evident, as is apoptotic cell death. The superficial endometrial layers become distended by the formation of hematomas; fissures subsequently develop, leading to the detachment of tissue fragments and the ultimate shedding of the functionalis. The resulting menstrual effluent contains fragments of tissue mixed with blood, liquefied by the fibrinolytic activity of the endometrium. Clots of varying sizes may be present if blood flow is excessive. The inflammatory components of menstruation may be essential for the rapid restoration of tissue integrity that occurs

following endometrial sloughing.171 The withdrawal of progesterone, as a well-­known antiinflammatory mediator of the secretory phase, likely participates in the onset of inflammatory changes, including induction of MMPs, urokinase-­ type plasminogen activator and tissue-­type plasminogen activator (uPA and tPA), and PAI1 expression.171–173 At menstruation and with progesterone withdrawal, prostaglandin-­endoperoxide synthase 2 cyclooxygenase-­2 (PTGS2 or COX-­2) is dramatically elevated via NFKB, with induction of prostaglandins and lipoxygenases (LOX).174 One LOX, LOX15, is expressed with progesterone withdrawal. Since LOX15 is responsible for production of the antiinflammatory eicosanoid lipoxin A4 (LXPA4), expression may help curtail inflammatory responses.175,176 Surprisingly, LXPA4 has also shown to be a potent estrogen receptor agonist177 and could function to facilitate endometrial repair, particularly during menstruation, when circulating estrogen levels are low. The duration of menses in ovulatory cycles is variable, generally 4 to 6 days, but usually similar from cycle to cycle in any individual ovulatory woman. The duration of flow is abnormal if it is less than 2 days or more than 7 days. The amount of blood lost in a normal menses ranges from 25 to 60 mL, and loss of more than 60 mL per month is associated with iron deficiency anemia. Stem Cells and Telomerase in Endometrial Renewal. Menstruation and cyclic repair of the endometrial lining require both stem cells and an adaptation involving telomerase that imbues the endometrium with near immortality, compared with other tissues in the body. Disorders of endometrial regeneration can result in severe pathological conditions, including uterine cancer, endometriosis, and infertility. Over the last two decades, different approaches have been used to identify putative stem cells in the human endometrium. To date, no consensus concerning endometrial stem cell markers has been established. However, studies in mouse and human have suggested that the endometrium possess both endogenous and exogenous sources of endometrial stem cells that contribute to the endometrium’s regenerative capability. In 2004, endometrium in women who had undergone bone marrow (BM) transplantation was examined and found to contain endometrial epithelial and stroma cells that had clearly been derived from the donor BM.178 This observation supported the idea that BM-­derived stem cells can traffic to and engraft in the endometrium, and secondly, that these undifferentiated BM stem cells undergo both mesenchymal and epithelial transformation adopting a phenotype indistinguishable from their host organ. This original observation was later confirmed in the mouse, using male BM donors and female recipients, proving that these were not endometrial stem cells that reside in the BM but rather BM-­ derived pluripotent stem cells. Endometrial-­derived pluripotent cells, whether native to the endometrium or derived from the BM, appear to have unusual properties that may revolutionize how we think about stem cell research.179 These cells have been manipulated in the mouse model to differentiate into neuronal cell types with the potential to produce dopamine, and pancreatic beta cells that produce insulin.180,181 Mesenchymal stem-­like cells that express CD140b and CD146 have also been identified that can be differentiated into adipocytes, osteocytes, chondrocytes, myocytes, and endothelium.182 Based on these early reports, the ready access to stem cells of endometrial origin could supplant the necessity to perform BM biopsy and offers the promise of a self-­renewing source of autologous stem cells for grafting of a wide variety of cell types that might cure such human diseases such as Parkinson or type I diabetes, as well as treatment for Asherman syndrome with the possibility of endometrial renewal.183–185 Similarly, several studies have suggested the regenerative potential of platelet-­ rich plasma (PRP). Treatment of human endometrial primary cells and in vivo animal models of Asherman syndrome with PRP leads to increased proliferation and regeneration of endometrial

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract

tissue.186–188 Although initial results are promising, further mechanistic and clinical studies are necessary.188–190 Early methods for identifying endometrial stem cells began with the study of retention of cells labeled with 5-­bromo-­2-­deoxyuridine (BrdU).154,191 These rare, highly proliferative and undifferentiated quiescent populations of cells maintain properties of clonicity, colony-­forming unit activity, and the ability to reconstitute their tissue of origin.146 Hoechst 33342 exclusion to identify the side population (SP) cells have been used to show that in the endometrium, these SP cells are maintained at a constant level (∼1%) throughout a woman’s reproductive life.146,155 Interestingly, it has been suggested that BM-­derived cells do not contribute to this population of SP cells in the endometrium, suggesting they are a resident to the endometrium and therefore provide an endogenous source of self-­renewing stem cells to the endometrium.192 Specific cell surface markers have been identified in endometrial mesenchymal stem cells (MSGs).179 These CD146+ PDGFRB− cells only represent less than 2% of endometrial stromal cells193 and are found in a perivascular location, similar to other BM-­derived cells. These cells have increased proliferation capacity and may be important in the reconstitution of the endometrium following menstruation. The endometrium, unlike other tissues in the body, does not appear to age. Despite up to 450 cycles of menstruation and regeneration, the endometrium of the typical woman continues to renew itself with remarkable reliability. Age is related to telomere length, which in other tissues determines the lifespan of a cell.194 Endometrium expresses the enzyme telomerase, at levels similar to certain cancers.195 Levels are cycle dependent, with higher levels in the proliferative phase reaching their nadir during the midsecretory phase.196,197 Interestingly, telomerase is found in the epithelial but not the stromal compartment.198 Epithelial stem cells are thought to participate in endometrial renewal following menses, arising from the basalis layer.199 These cells appear to express stage-­specific embryonic antigen 1 (SSEA-­1 or CD15), a Lewis X epitope found on embryonic stem cells. The endometrial cells expressing this epitope had greater telomerase activity than cells without the epitope. Interestingly, telomerase may be indirectly regulated by estrogen, through WNT pathways involving CTNNB1 expression.200 In women and baboons with endometriosis and progesterone resistance, telomerase levels appear to be increased, suggesting a possible link to this disease and the pathophysiology of its chronicity and tendency to recur.201 Vascular Remodeling and Angiogenesis. Angiogenesis, the formation of new blood vessels from preexisting vessels, rarely occurs without injury or disease in the normal adult, except in the female reproductive tract and ovary. Here the cyclic processes of endometrial shedding and regeneration and corpus luteum formation entail remarkable changes in vessel growth and remodeling. The angiogenic process involves multiple steps and is tightly regulated by activators and inhibitors.202–204 There are four phases of the endometrial cycle when important events relating to angiogenesis occur: (1) at menstruation, when there is a repair of ruptured blood vessels; (2) during the proliferative phase, when there is a rapid growth of endometrial tissue; (3) during the secretory phase, with the development of the spiral arterioles that feed a subepithelial capillary plexus; and (4) in the premenstrual phase, when there is evidence for vascular regression. If this angiogenic remodeling program is not properly executed, abnormalities in endometrial function can result, including menorrhagia. Angiogenesis during the proliferative phase is by vessel elongation.205 In the secretory phase, intussusception appears to account for the increase in vessel branching; this proliferation of endothelial cells inside vessels ultimately produces a wide lumen that can be divided by transcapillary pillars or alternatively lead to capillary fusion or splitting. Although most prominent in the

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late menstrual and early and late proliferative phase, endothelial cell proliferation is continuous during the menstrual cycle. Thus vessel growth continues during the secretory phase, despite the fact that the surrounding endometrial tissue has ceased to grow, resulting in the coiling of spiral arterioles. Endometrial angiogenesis and vessel remodeling are directed by a network of signaling molecules and receptors that include members of the vascular endothelial growth factor family, FGFs, angiopoietins, angiogenin, and the ephrins and their cognate receptors—which can also exist as secreted ligand-­binding domains that function as inhibitors as a result of alternative splicing.202–204,206 Although the temporal and spatial patterns of expression of several other angiogenic factors and their receptors have been defined in the endometrium, the specific roles of each of these factors in the endometrial angiogenesis–vessel remodeling cycle remain to be elucidated. Of the members of the vascular endothelial growth factor family that includes VEGF-­A, VEGF-­B, VEGF-­C, and VEGF-­D, VEGF-­A is most important for endometrial angiogenesis.205–207 VEGF-­ A acts on two different receptors: VEGF receptor-­ 2 (VEGFR2), which may play the dominant role in signaling endothelial cell proliferation; and another tyrosine kinase receptor, VEGFR1 (also known as FLT-­1), which may play the dominant role in mediating VEGF effects on vascular permeability.208–211 Both receptors are present on endothelial cells. VEGFR2, also known as kinase domain receptor (KDR), has been detected in stromal and epithelial cells of the premenstrual endometrium; this presence suggests actions on nonvascular compartments. The premenstrual phase is characterized by a dramatic upregulation of the VEGFR2 receptor in stromal cells of the superficial layers of the endometrium in response to progesterone withdrawal. The action of VEGF on VEGFR2 may participate in the increased expression of MMP-­1 in the stromal in the premenstrual phase. VEGF-­A expression is detectable in glandular epithelial and stromal cells in the proliferative phase, stimulated by estrogen through the actions of hypoxia-­ induced factor-­ 1 alpha (HIF1A).212,213 HIF1A is also induced by prostaglandins E2, which is maximally produced at the time of menstruation.214 VEGF released from neutrophils in intimate contact with the endothelial cells is thought to stimulate endometrial vessel growth. It is also present in uterine NK cells. During the secretory phase, VEGF-­A can be identified in surface epithelial cells, which presumably secrete into the uterine cavity. The release of VEGF from subepithelial NK cells has been suggested to play a role in directing the development of the subepithelial capillary network in the secretory phase. VEGF-­A has four common isoforms. It has four common splice variants (VEGF121, VEGF165, VEGF189, and VEGF 206). After ovulation, there is a remarkable shift in VEGF-­A isoforms expressed in the uterus with the appearance of VEGF-­A189 in the perivascular stromal cells; the VEGF-­A189 isoform can be processed by proteolytic cleavage by plasminogen activator.113 VEGF-­A189 increases vascular permeability acting on VEGFR1, while its processed form binds to the VEGFR2 receptor, which mediates the mitogenic action on endothelial cells. The highest VEGF-­A levels are found in the menstrual phase, probably a response to proinflammatory cytokines. The surge might also be attributed to focal hypoxia, which is a potent stimulus to VEGF-­A gene transcription. Expression of VEGFR1 and VEGFR2 is also greatest in the menstrual phase. The elevated levels of VEGF and cognate receptor expression at this time are presumed to be important for vessel repair and the preparation for angiogenesis in the proliferative phase. Notably, the functional activity of VEGFR2 (as assessed by receptor phosphorylation), a signature indicating ligand activation, is relatively low in the early menstrual phase when levels of soluble VEGFR1 (sFLT-­ 1), which sequesters VEGF, are highest. VEGFR2 receptor

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phosphorylation increases substantially in the late menstrual phase, remaining modestly elevated in the early and late proliferative phase when sFLT-­1 levels decline.215 The FGF family of proteins may also participate in endometrial angiogenesis through interactions with the VEGF system. FGFs upregulate VEGFR2 and VEGF-­A expression, and in a feedforward loop, VEGF-­A promotes the release of FGFs from the extracellular matrix. Basic FGF is a potent stimulus for ανβ3 integrin expression. This integrin is present at the site of active angiogenesis and fundamental to endothelial invasion and vessel elongation during angiogenesis.216 The angiopoietins (Ang) regulate vessel stability. Ang-­ 1, expressed in vascular smooth muscle cells, binds to a cognate receptor, Tie-­2, on endothelial cells, resulting in vessel stabilization. Ang-­2 is a physiologic antagonist of Ang-­1. It also binds to the same Tie-­2 receptor. Vessels atrophy when Ang-­2 acts in the absence of VEGF-­A, whereas angiogenesis is promoted in the presence of VEGF-­A. In situ hybridization studies indicate that Ang-­1 expression is most abundant in the glands and stroma of the early and midproliferative phase and reduced in the late proliferative phase.217 Ang-­2 expression is detected in the glands and stroma throughout the cycle, with the highest expression occurring in the early proliferative and mid-­to late secretory phases. In endometrium from women with menorrhagia, Ang-­1 expression is consistently downregulated; as a result, the ratio of Ang-­1 to Ang-­2 is reduced, which contributes to vessel instability.217 Angiogenin is a heparin-­binding molecule that is expressed by endometrial epithelial and stromal cells at the greatest levels in the mid-­to late secretory phases and the decidua of early pregnancy. Angiogenin is thought to contribute to the proliferation of vascular smooth muscle cells around the spiral arterioles. Like VEGF-­A, expression of angiogenin is stimulated by hypoxia. It is also increased by progesterone. Ephrins, a family of molecules and their cognate tyrosine kinase receptors, are believed to guide endothelial cells to specific targets. Ephrins have been detected in endometrial endothelial and stromal cells, but the functional roles of these molecules and their receptors in the uterus remain to be clarified. The physiologic consequences of angiogenesis are reflected in changes in endometrial blood flow. By measuring the clearance of radioactive xenon gas, the highest endometrial perfusion was reported between days 10 and 12 and days 21 and 26 of the cycle.94 Microvascular perfusion has been assessed by laser Doppler fluximetry with transvaginal placement of a fiberoptic probe into the uterine cavity.218 With the use of this technique, endometrial perfusion was found to be highest during the proliferative phase and the early secretory phase, not too dissimilar from the finding based on xenon clearance. Uterine blood flow

is greatest in the fundus, and higher flow rates are associated with better outcomes in assisted reproduction. Notably, diminished uterine blood flow has not been found in the perimenstrual period, but these methods cannot easily identify localized areas of vasoconstriction. Extracellular Matrix Remodeling. The biochemical basis for the dramatic structural changes in the endometrium in the perimenstrual period includes the action of specific matrix-­ degrading proteases, the MMPs.158,162–165 Studies on human endometrial explants in culture demonstrated that degradation of the extracellular matrix occurs in the absence of progesterone and estrogen, which suppress the expression MMPs. Moreover, this degradative process can be blocked by MMP inhibitors but not by inhibitors of lysosomal cysteine proteinases—directly implicating MMPs in the catabolism of the endometrial extracellular matrix. Enzymes of the fibrinolytic system (urokinase and tissue plasminogen activators) are increased in the endometrium as progesterone is withdrawn in the perimenstrual period. Moreover, PAIl expression is reduced, allowing the plasminogen activators to activate plasmin and proteolytically cleave and activate the latent MMP proenzymes.219 MMPs represent a large family of proteinases that play a major role in the remodeling of the extracellular matrix (Fig. 10.9). In situ hybridization and immunocytochemistry have been used to map the expression of MMPs and the endogenous inhibitors, tissue inhibitors of the matrix metalloproteinases (TIMPs), in the primate endometrium. Cell-­ specific and menstrual cycle-­specific patterns were revealed, with the most profound changes occurring during the perimenstrual period.163,220 After ovulation, the expression of interstitial collagenase (MMP-­1), stromelysin-­1 (MMP-­3), and stromelysin-­2 (MMP-­10) in the endometrial stroma is essentially restricted to the perimenstrual and menstrual phase. Other MMPs are detected during the proliferative and secretory phases but are significantly increased in expression perimenstrually. These include the type IV collagen-­degrading enzymes, MMP-­2 and MMP-­9. The membrane-­bound MMP, MMP-­14 (which activates MMP-­2), is detected during menstruation in stromal inflammatory cells and epithelial cells. TIMP-­1, which is detectable in the endometrium throughout the cycle, is increased in the stroma, epithelium, and arterioles at menstruation. The importance of progesterone withdrawal in regulating endometrial MMPs and the different temporal patterns of expression have been well-­documented in in vivo and in vitro systems.104 In a primate model in which hormone levels were manipulated by steroid implants, progesterone withdrawal resulted in

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upregulation of MMP-­1, -­2, -­3, -­7, -­10, -­11, and -­14. It is important to note that the expression of MMPs in the endometrium is heterogeneous.158,220 At the start of menstruation, MMP-­1 is found in patches of stromal cells in the superficial zone; these patches are colocalized with areas of reduced stromal and epithelial expression of estrogen and PR and focal disruption of the extracellular argyrophilic fibrillar network—reflecting the degradative activity of MMP-­1. As the process of menstruation proceeds, MMP-­1 expression spreads to include the entire functionalis. MMP-­2 and MMP-­3 expression is also limited to the stromal cells in the functionalis. During menses, MMP-­1, -­2, -­3, and -­9 localize primarily in and around arteriolar walls. The heterogeneity of MMP expression suggests that MMP gene transcription is under the control of local rather than systemic (steroidal) factors. In other words, steroids are indirectly influencing MMP expression. Progesterone, particularly in the presence of estradiol, can suppress the expression of certain MMPs (i.e., MMP-­1, -­2, -­7, -­9, and -­11) in endometrial explant culture.165 This action is most likely explained by changes in autocrine/paracrine signals—particularly proinflammatory cytokines or members of the transforming growth factor family, which respectively are potent inducers and suppressors of MMP gene transcription. In culture systems, IL-­lα has been implicated as the mediator of MMP-­1, MMP-­3, and MMP-­7 expression in response to withdrawal of progesterone. Neutralizing antibodies to TGF-­ β prevents the action of progesterone in blocking MMP-­3 and MMP-­7 expression. EBAF is the orthologue of the murine gene named Lefty and another likely candidate for a progesterone-­regulated cytokine controlling MMP expression.221–223 LEFTY was originally identified in human endometrium as a gene upregulated in the late secretory and menstrual phases of the normal cycle, being absent in the proliferative, early, and midsecretory endometrium. EBAF expression, which is predominantly found in the endometrial stroma and to a much lesser extent in the glandular epithelium, is suppressed by progesterone. Interestingly, endometrium from women with a history of abnormal bleeding and endometriosis expressed Lefty at unusual times including the proliferative, early, and midsecretory phases.224 Unlike other members of the TGF-­β family that promote the formation and stability of the extracellular matrix, EBAF downregulates the elaboration of collagen in association with reduced expression of connective tissue growth factors while upregulating expression of collagenolytic and elastinolytic enzymes.225 These actions of EBAF are the result of antagonism of the SMAD signaling pathway that is activated by the other TGF-­β growth factors. Thus the decline in progesterone and estradiol in the late luteal phase initiates alterations in the endometrium that include upregulation of proinflammatory cytokines (some of which may be contributed by immune cells that accumulate in the endometrium) and a natural TGF-­β antagonist. The collective result is focal and widespread expression of matrix-­degrading enzymes that result in the remodeling of stroma and blood vessels in the functionalis. Lysosomal involvement in the process of menstruation has been proposed because of three observations: an increase in the abundance of lysosomes in the endometrium during the late secretory phase, the cytochemical demonstration of acid phosphatase in the perimenstrual endometrium, and the high specific activity of certain lysosomal hydrolases in endometrial tissue in the menstrual phase.226 However, inhibitors of these enzymes, leupeptin and E-­64, do not prevent the progesterone withdrawal-­induced breakdown of extracellular matrix in endometrial explants, as do the inhibitors of MMP activity. These observations suggest that lysosomal proteinases are not major contributors to the remodel ing of the perimenstrual endometrium.

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Fig. 10.10 Changes in PGF2α levels and endometrial 15-­hydroxy prostaglandin dehydrogenase (PGDH) activity. (From Casey ML, Hemsell DL, MacDonald PC, Johnston JM. NAD+ dependent 15-­hydroxy prostaglandin dehydrogenase activity in human endometrium. Prostaglandins. 1980;19:115; and Demers LM, Halbert DR, Jones DF, Fontana J. Prostaglandin F levels in endometrial jet wash specimens during the normal human menstrual cycle. Prostaglandins. 1975;10:1065.)

Vasoactive Substances. The endothelins are a family of potent vasoconstrictors produced by endothelial cells that act on two types of receptors present on vascular smooth muscle. Endothelin-­ 1, produced by endometrial epithelial or stromal cells, may act on spiral artery smooth muscle cells to promote vasoconstriction. Enkephalinase, a membrane-­ bound metalloendopeptidase, degrades endothelin-­1 and other vasoactive peptides and is present at the highest levels in the midsecretory endometrium.129 Expression of the gene encoding enkephalinase is upregulated by progesterone. The decline in progesterone levels at the end of the luteal phase results in a subsequent fall in enkephalinase, which prolongs the biological life of endothelin-­ 1. Vasopressin may also function as a vasoconstrictor in the endometrium during the menstrual phase of the cycle.227 The production of prostaglandins, particularly PGF2α and other eicosanoids in the endometrium, is enhanced by lysosomal phospholipases that liberate the arachidonic acid that accumulates in the endometrium during the secretory phase; in turn, arachidonic acid is metabolized into prostanoids (see Chapter 4).228 The premenstrual decrease in progesterone is also followed by induction of the prostaglandin synthase, COX-­2, and a decline in 15-­hydroxyprostaglandin dehydrogenase activity, which inactivates PGF2α. This induction of the prostaglandin synthase, COX-­2, and the decline in 15-­hydroxyprostaglandin dehydrogenase leads to increased production and bioavailability of PGF2α, which triggers myometrial contractions that compress the endometrial vasculature and promote hemostasis (Fig. 10.10).229 Increased PGE2 coupled with hypoxia stimulates IL8 expression with a role in endometrial repair.230 Hemostatic and Fibrinolytic Mechanisms. The relative activities of the hemostatic and fibrinolytic systems in the endometrium are shifted in the perimenstrual period such that clotting activity is reduced and fibrinolytic activity is increased. Consequently, menstrual fluid does not normally clot, even during prolonged storage. Decidualized stromal cells express TF, the primary trigger of thrombin formation and hemostasis, under the influence of progesterone.231 Expression of TF is expressed in the endometrial stromal cells along with another hemostatic factor, PAI-­1, under the influence of progestins, which suppresses MMP production.232 TF production by the decidualized stromal cells declines with the withdrawal of progesterone, along with the

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The endometrial fibrinolytic system includes uPA and tPA, which cleave plasminogen to yield the fibrinolytic enzyme, plasmin.231,233,234 Progesterone reduces expression of urokinase and increases that of PAI1 in cultured endometrial cells. Removal of progesterone or the addition of the antiprogestin, RU486, reverses these responses.234

ENDOMETRIUM IN THE CYCLE OF CONCEPTION AND PREGNANCY Secreted Proteins of the Endometrium In preparation for pregnancy, the endometrium produces a large number of secreted proteins that are collectively referred to as the endometrial secretome.235–237 These proteins serve critical autocrine, paracrine, and juxtacrine roles for the developing endometrium and embryo.237–240 The uterine glandular epithelium secretes factors such as glycogen that are thought to serve as a histotrophic nutrition source for the embryo before 10 weeks’ gestation.241 In addition to macromolecules, the endometrium also is likely to selectively secrete many small molecules, since it is well endowed with many members of the adenosine triphosphate-­ binding cassette transporter protein family, which are involved in the regulated secretion of a variety of small molecules, including drugs, lipids, and conjugated molecules.242 Endometrial glandular secretions provide early nourishment for the embryo and signal the invading trophoblast.237,240 A vascular circulation to the embryo does not occur until the end of the first trimester, making these uterine secretions of critical importance. The vital role of uterine gland-­derived factors in the establishment of pregnancy has been demonstrated for large animals and mice but is only implied in humans.243 The human endometrium has a high density of uterine glands, approximately 15 glandular openings per mm2, and an increase in glandular area is observed in the secretory phase compared to the proliferative phase.244,245 Women undergoing ART and receiving clomiphene citrate have significantly reduced glandular area compared to stage-­matched controls.244 Thus, it is proposed that defects in uterine gland development or function are indicative of a suboptimal endometrial environment. Endometrial secretory proteins can be detected in the lumen of the uterine cavity and some are found in the general circulation. A remarkable increase in secretory activity is associated with the luteal phase and early pregnancy, primarily in the glandular epithelial cells and later the decidua. These secreted proteins are either directly or indirectly regulated by progesterone, and most are implicated in the regulation of embryo implantation, trophoblast invasion, and/or early embryo survival.246–248 Therefore an understanding of the mechanisms of progesterone action allows significant insight into endometrial function.

Glycogen Glycodelin, the most abundant product of the secretory phase endometrial glands and decidua, is known by several names: progesterone-­associated endometrial protein, α-­uterine protein, pregnancy-­ associated endometrial alpha 2 globulin, endometrial protein 15, chorionic alpha2 macroglobulin, and placental protein 14 (an erroneous designation because glycodelin is an endometrial/decidual protein).248,249 The mature form of glycodelin contains 162 amino acid residues and is 17.5% carbohydrate by weight. It has extensive structural homology with the beta-­lactoglobulins and, to a lesser extent, with retinol-­binding proteins. Glycodelin is a glycoprotein with four isoforms.249 Glycodelin A is the major progesterone-regulated protein secreted into the uterine lumen. However, its functions with

respect to the endometrium, implantation, and pregnancy are still largely unknown. Glycodelin A is a potent inhibitor of sperm-­egg binding, and researchers have postulated that it may play a role as an immunomodulator because of its ability to suppress NK cells.250,251 Glycodelin has been shown to limit trophoblast invasion in vitro by suppressing MMP production by cytotrophoblast.252 The absence of glycodelin A during the time of fertilization and its appearance during the time of implantation and placentation are consistent with these proposed functions.253 Glycodelin levels in uterine flushings are tightly correlated with the histologic date of the endometrium (Fig. 10.11). It is not detectable in significant amounts in uterine flushings during the follicular and early luteal phases. Six days after ovulation, however, the levels rise rapidly to reach a concentration 100 times higher than plasma levels. In peripheral serum, glycodelin appears 5 days after ovulation, reaching peak concentrations in nonfertile cycles at about the time of menstruation; levels reach a nadir during the midfollicular phase of the subsequent cycle. In a cycle of conception, glycodelin levels increase rapidly after implantation, reaching a maximum at 8 to 10 weeks of gestation and subsequently declining in a pattern that mimics the changes in hCG. The discordant pattern of glycodelin and progesterone in serum may reflect the slower turnover of the protein. Levels of glycodelin in serum fail to rise in women using combination-­ type oral contraceptives and in some patients with luteal-­phase defects.254 There is a good, but not perfect, correlation between the progestational activity of steroids and their ability to stimulate endometrial glycodelin synthesis. Relaxin has also been implicated as a stimulus for glycodelin expression.253,255 Specific histone deacetylase inhibitors have been shown to potentiate the action of progesterone on both endometrial epithelium and stroma.256 The histone deacetylase inhibitor trichostatin A (TSA) has been shown to induce glycodelin in Ishikawa cells. Further, enhanced placental cell (JAR) spheroid attachment to Ishikawa cells was found by upregulating glycodelin using an in vitro model of implantation.257 This and other studies suggest a role for glycodelin in both the differentiation and function of the receptive endometrium. Glycodelin has been shown to be decreased in the endometrium of women with infertility, including those with luteal phase defect254 and PCOS.258 In women exposed to the levonorgestrel-­ containing intrauterine device (IUD) or the Yuzpe emergency contraception regimen, glycodelin expression was unaffected or increased.259,260

Insulin-­Like Growth Factor Binding-­Protein One Insulin-­like growth factor binding-­protein one (IGFBP-­1), also known as pregnancy-­associated alpha1 protein and placental protein 12, is a major secretory product of decidual cells.261 It is one of several proteins that bind IGF-­1 and IGF-­2, affecting the ability of these growth factors to interact with the IGF receptors. Consequently, the binding proteins can have significant roles in modulating IGF effects. The protein undergoes posttranslational modification by phosphorylation, which increases its affinity for IGF-­1 and therefore its ability to neutralize IGF-­1 action. IGFBP-­ 1 derived from the decidua has been proposed to control the invasion and proliferation of trophoblast cells during implantation and placentation by sequestering IGFs. In a transgenic mouse model, overexpression of IGFBP-­1 in the decidua resulted in abnormal placental morphology due to defects in trophoblast invasion and differentiation.262 Because IGFBP-­1 contains the Arg-­Gly-­Asp (RGD) motif recognized by cell surface integrins that bind fibronectin, its actions may be more complex than simple IGF sequestration when it is presented to cells that

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termination with the antiprogestin RU486 causes a marked decline in IGFBP-­1 levels before a decrease in hCG levels, confirming the progesterone dependence of IGFBP-­1 production by decidual cells. Defects in IGFBP-­ 1 have been noted in certain conditions associated with infertility and pregnancy loss, including PCOS and endometriosis.258,270 Such observations may reflect an altered endocrine milieu, higher insulin levels, or a relative progesterone resistance in these conditions (discussed in subsequent text). In vitro, insulin inhibits the normal process of endometrial stromal differentiation (decidualization). In addition, hyperinsulinemia downregulates hepatic IGFBP-­1, resulting in elevated free IGF-­I in the circulation. Thus the elevated androgens and estrogen seen in PCOS, along with decreased progesterone in the absence of ovulation, likely contribute to endometrial dysfunction, infertility, increased miscarriage rate, endometrial hyperplasia, and endometrial cancer common in women with PCOS.

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Fig. 10.11 Glycodelin levels in endometrium, serum, and amniotic fluid. (Reprinted with permission from Seppala M, Koistinen H, Koistinen R. Glycodelins. Trends Endocrinol Metab. 2001;72:111–117.)

Stromal cell IGFBP-­1 mRNA levels are regulated by progesterone as well as by hypoxia, IGFs, insulin, relaxin, and other growth factors.264–266 Both IGF and insulin decrease decidual IGFBP-­1 release, while relaxin increases its release in a dose-­dependent manner. HOXA10 may have a mild impact on IGFBP-­1 expression, but in the presence of the forkhead/ winged-­helix transcription factor, FOX01, IGFBP-­1 is markedly stimulated.267 FOX01, prolactin, and IGFBP1 are all induced during decidualization and likely are involved in the progesterone-­mediated stromal differentiation and prevention of apoptosis in preparation for nidation. Intrauterine microdialysis studies revealed that IGFBP-­ 1 is released into the uterine lumen in the late secretory phase (10 days after ovulation or later)268 and increased secretion in response to hCG.268 IGFBP-­1 is increased between midsecretory to 6 weeks of pregnancy, as demonstrated by DNA microarray analysis on endometrium from ectopic pregnancy.269 IGFBP-­1 levels then decline in the second trimester only to rise again in late pregnancy, also accumulating in amniotic fluid. Pregnancy

Osteopontin (OPN) is a member of the SIBLING family of proteins.271,272 Each of the proteins in this large family contains the three amino acid sequence Arg-­Gly-­Asp (RGD) and has binding sites specific for two major cell surface receptors, ανβ3 and CD44.271 OPN is a 70-­kDa glycosylated phosphoprotein secreted by the glandular epithelium and is expressed during the midsecretory phase, localized to the luminal endometrial epithelium.273 OPN is regulated by progesterone.273,274 The secretion of OPN and subsequent binding to the luminal surface suggests direct interaction is occurring between integrins and this molecule, with a role as a “sandwich” ligand that serves as a bridge between surface receptors on the endometrial and embryonal surfaces. Alternative roles for OPN and these receptors include the prevention of complement fixation as part of a protective mechanism involving the innate immune system.275

Prolactin Prolactin is produced by both the endometrium and the myometrium.276,277 Serum levels of prolactin do not change across the menstrual cycle but rise markedly during the first trimester of pregnancy in parallel with the decidual response, reaching peak concentrations at 15 to 20 weeks of gestation. Prolactin, like IGFBP-­1, is a biomarker of human endometrial stromal cell decidualization. Regulators of decidualization include ovarian steroids, cAMP or forskolin, IL-­11, and PGE2. Recent studies suggest upstream mediators of decidualization may regulate prolactin expression, including various proteins expressed during the secretory phase, including ghrelin, IGFBP-­related protein-­1, leptin, and oncostatin-­M.278–282 The proliferation of endometrial cells is inhibited during decidualization, perhaps through stromal factors, such as IL-­6 and oncostatin-­M.278,283 The role of PRL in the decidualization process remains unclear. Prolactin receptor (PRL-­ R) is expressed on the endometrial stroma, and while PRL-­R knockout mice had defects in decidualization, the phenotype included reduced expression of LIF and a number of other factors that could be mitigated by exogenous progesterone supplementation. This suggests that ovarian PRL-­R stimulation by prolactin inducing progesterone secretion is a critical event in mice, rather than any direct effect of prolactin acting directly on the decidualizing stroma.284 A role for uterine prolactin controlling the NK cells has recently been suggested.285 Prolactin produced by the decidua during pregnancy accumulates in the amniotic fluid, where it has been postulated to have effects on osmoregulation and fetal lung

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A

Fig. 10.12 Proliferative (A, prereceptive) and midsecretory (B, receptive) endometrium is regulated by ovarian steroids, resulting in successful implantation of the human embryo (C). (Generously provided by Dr. Allen Enders, from the Carnegie Collection of early human implantation sites. Used with permission from Human Reproduction.)

B

C

Endometrial Preparation for Implantation The coordinated actions of ovarian estrogen and progesterone coordinated endometrial preparation for implantation. After ovulation, rising progesterone from the nascent corpus luteum transforms the proliferative endometrium into a secretory structure receptive to embryo implantation (Fig. 10.12A and B; see Chapter 7).121,246 Uterine transformation is required to ensure that the growing embryo will interact with the surface epithelium and invade the underlying stroma (Fig. 10.12C). Implantation can be divided into distinct and separate stages, coordinated primarily by estrogen and progesterone action on the female reproductive tract (Fig. 10.13).286 Following ovulation, the ovum enters the fallopian tube, where fertilization occurs. The early cell divisions ensue and the embryo enters the uterine cavity at the morula stage, approximately 3 to 4 days after being fertilized. Implantation begins about the sixth to seventh day after fertilization. The initial attachment reaction (apposition) may be the rate-­limiting step, and failure to adhere may preclude the subsequent stages of implantation. Studies in experimental and domestic animals have demonstrated the synchronous development of the embryo and endometrium that is required for normal implantation and development to occur.238,287–290 In laboratory animals and women, there is a discrete “window” of time for implantation, which in some species lasts for only a matter of hours.71,238,289,291–295 In women, the period of receptivity is known as the “window of implantation” and begins on day 19 or 20 of the cycle and remains open for 4-­5 days (LH+5 to +9) as progesterone reaches peak serum concentrations.296

Uterine receptivity is defined as the period of endometrial maturation during which the trophectoderm of the blastocyst can attach to the endometrial epithelial cells and subsequently proceed to invade the endometrial stroma to establish a healthy pregnancy. Indeed, blastocyst implantation outside of that window results in a higher miscarriage rate, possibly due to a shift in loss of synchrony between endometrium and embryo (Fig. 10.14). This explanation may provide insights into some cases of otherwise unexplained infertility (UI) and unexplained recurrent pregnancy loss (RPL). A functional endometrium and the timing of embryo implantation are central to a successful pregnancy, yet our current clinical assessment relies heavily on morphologic features (endometrial thickness and pinopods) that provide little significant clinical information in human pregnancy. Identification of molecular markers predictive of a functional endometrium capable of supporting pregnancy is an important goal to treat infertility, increase ART pregnancy rates and decrease pregnancy-­associated health complications resulting in diminished fetal survival and maternal health. Classically, the assessment of endometrial receptivity has relied mainly on morphologic features (endometrial thickness and pinopods) and current research has focused on specific biomarkers to assess the endometrial status and predict pregnancy success, including growth factors, lipids, cytokines, and adhesion molecules.297 These studies have not identified any molecular or histological marker that can be reliably used to define that state of endometrial receptivity in clinical practice. Thus, embryo transfer in ART clinics is dictated solely by the quality of the embryo and the thickness of the endometrium. With the

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Fig. 10.13 Stages of implantation beginning with ovulation, fertilization, then transport to the waiting endometrium. Implantation depends on synchronous development between the endometrium and the embryo.

100 Embryo implantation Miscarriage

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Days After LH Surge Fig. 10.14 The timing of implantation in a population of fertile women.  Most successful pregnancies implant between 7 and 10 days following a urinary luteinizing hormone (LH) surge (blue line), but the probability of miscarriage increases when implantation occurs outside of this 4-­day window (red line). (Modified from Wilcox A, Barid DD, Winberg CR. Time of implantation of the conceptus and loss of pregnancy. N Engl J Med. 1999;240:1796–1799.)

advent of the endometrial receptivity array (ERA), individual receptivity status can potentially be assessed. Developed in 2011, the ERA analyzes 238 genes and with a computational algorithm detects the unique transcriptomic signature of endometrial receptivity.298 Of note, personalized embryo transfer based on the ERA in women with recurrent implantation failure resulted in implantation rates comparable to normal receptive controls.299 The unique transcriptomic signature of endometrial receptivity provides potential therapeutic targets for the treatment of infertility and alternatively the development of nonhormonal contraceptives.

Early Implantation Events Implantation can be viewed as a highly complex and orchestrated interaction between the maternal endometrium and the newly formed embryo.239,286 As depicted in Fig. 10.15, multiple soluble and membrane-­bound factors have been elucidated that facilitate embryo growth, differentiation, attachment, invasion, and avoidance of immunologic rejection. Maternal factors appear to simultaneously permit intrusion while limiting the degree of embryonic invasion into maternal tissue. Many of the embryonic signals or receptors have complementary ligands or

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Villi

Blastocyst hCG, MMPs, IL-1 uPA, TNF , PDGF EGF rec, integrins, fibronectin, HLA-G, trophinin, CD44 Integrins, HB-EGF, OPN CD44, pinopodes, mucins

Trophoblast MMPs, integrins, uPA and receptor, IL-1, TGF , HLA-G

LIF, calcitonin, glycodelin, OPN

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VEGF, fibronectin, HB-EGF, TGF , integrins, GM-CSF, KGF, EGF, PGs, IL-11 LIF, IGFBP1 Vasculature

Leukocytes IL-1, IL-2, TNF , INF , bFGF

coreceptors on the maternal surface. Mimicking maternal antigens by the invading embryo is a strategy that is also used by the embryo to penetrate the endometrium, without triggering host defenses.300 In the event of oocyte fertilization, hCG from the implanting embryo rescues the corpus luteum from apoptosis and regression, maintaining serum progesterone secretion and endometrial integrity. This integrated mechanism, demonstrating the importance of synchrony, creates a window of endometrial and ovarian receptivity that is essential for the success of the pregnancy.293 Women who are delayed in establishing embryo implantation, beyond the normal window of implantation (7 to 10 days after ovulation), are at increased risk for miscarriage. The examination of hysterectomy specimens revealed that the first consistent structural changes in the endometrium of early pregnancy are recrudescence or accentuation of glandular secretory activity, edema, and the predecidual reaction.301 The increased prominence of the vasculature is considered to be a manifestation of increased blood flow, which together with VEGF production (a vascular permeability factor) may account for the associated edema. Endometrial biopsies in a cycle of conception suggest that stromal edema and vascular congestion are the earliest persistent morphologic features of the endometrium of pregnancy.302 Within the first weeks of gestation, the endometrium undergoes characteristic changes, in which epithelial cells of the endometrial glands become distended with clear cytoplasm. Many of the epithelial cells develop enlarged and hyperchromatic nuclei. The enlarged nuclei are polyploid. Parallel channels of the endoplasmic reticulum and large mitochondria are abundant in the epithelial cells, and the Golgi complexes have numerous stacked saccules. These changes are commonly referred to as the Arias-­Stella reaction 303 304 The ultrastructural characteristics of the endometrium are consistent with a hypersecretory state,

Fig. 10.15 Schematic showing some of the factors and their cells of origin during normal implantation in women.

providing the earliest nutrition and maternal communication with the nascent embryo. As blood flow to the placenta requires vascular remodeling that occurs later, the glandular histotroph provides direct and essential sustenance for the early fetus.305 Following the initial attachment reaction, the embryo invades the epithelium and the underlying decidualizing endometrial stroma. Decidualization is characterized by coordinated proliferation and differentiation of endometrial stromal cells into large epithelioid decidual cells, the secretory transformation of the uterine glands, an influx of specialized uterine natural killer cells, and vascular remodeling.306,307,308,309,310, The decidualized stroma represents a tissue that is both permissive and simultaneously a restrictive barrier to trophoblast invasion and placentation; its remodeling is crucial to the morphogenesis of the placenta and the establishment of the uteroplacental circulation. Moreover, the decidualized stroma represents the arena where the fetal semiallograft is exposed to maternal immunologically competent cells. While creating a hospitable environment for trophoblast invasion, the decidua also sets limits on this process to prevent excessive penetration and tissue destruction beyond its bounds. The polygonal decidual cells are arranged in a cobblestone configuration. The ultrastructural features of the decidual cells— including prominent Golgi complexes, dilated rough endoplasmic reticulum, and dense membrane-­bound secretory granules—are characteristic of secretory cells. The histologically distinct cell borders around decidual cells reflect the accumulation of a pericellular matrix (Fig. 10.16).184,185 Abundant decidual cell prolyl hydroxylase, an enzyme involved in collagen synthesis, indicates the important role of these cells in extracellular matrix production. There is also an abundance of amorphous components, including high-­molecular-­weight proteins with voluminous saccharide moieties (e.g., heparin sulfate proteoglycan). Fibrillar collagens are partially broken down and reorganized. Type V collagen epitopes are unmasked; collagen type VI, “stiff” short

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10

Fig. 10.16 Decidua of pregnancy.  Decidualized stromal cells are plump and have distinct cell borders. Glands are atrophic.

collagen fibers that bridge other fibrillar collagens disappear from most of the stroma and persist only in association with vessels and the basement membrane of the glands. The deposition of a basement membrane-­type matrix containing laminin and type II collagens around the decidual cells contributes to the formation of a “looser stroma” that serves as a substrate for the invading trophoblast cells. For example, entactin, a component of this basement membrane-­like matrix, promotes trophoblast cell adhesion and migration. The decidual matrix is also a rich source of cytokines, protease inhibitors, protease precursors, and other factors that modulate cell behavior. These are derived, at least in part, from the decidual cells whose secretory products also include IGFBP1 and TGF-­β, which may restrain the invasion of trophoblast cells.274,275 FOXO1A has been shown to regulate many of the genes induced during decidualization including IGFPB-­1 and prolactin.311 Ghrelin is an endogenous ligand for the growth hormone (GH) secretagogue receptor and has also been shown to be involved in the decidualization of human endometrium as well.277,278 Ghrelin, in combination with sex steroids estradiol and progesterone, and cAMP, increase both prolactin and IGFBP-­1, while ghrelin alone antagonizes the actions of cAMP.312 Reduced ghrelin has been associated with infertility.313 Prior to interaction with the surface epithelium, the blastocyst must hatch from the confines of the zona pellucida. Gradual zona thinning, as well as complete hatching of embryos, can be observed in vitro. The existence of ectopic implantation suggests that the endometrium is not obligatory for this process to be successfully completed. Nevertheless, there may be more subtle regulation of hatching within the endometrial cavity. Although degradation of the zona pellucida is a process controlled by the embryo, inhibitors or agents that induce “zona hardening” may affect the timing of the process. Work on nonhuman primates demonstrates that mononuclear cytotrophoblasts of the trophectoderm of the blastocyst have fused

into syncytia before the attachment of these cells to the endometrial epithelium.314 Careful histologic descriptions of very early human implantation sites (such as those studied by Hertig et al301) indicate that the syncytial trophoblast layer of the human embryo comprises the invading front during the first few days of implantation. Thus, it appears to be that it is a syncytial trophoblast cell that initially interacts with and adheres to the endometrial epithelium; only after the human embryo is completely embedded in the endometrium do cytotrophoblast cell columns start to stream out of the trophoblastic shell and further invade the uterus.315,316 This process starts approximately 1 week after the initiation of implantation and continues well into the second trimester of pregnancy. There is growing consensus that initial apposition and attachment is an evanescent and rate-­limiting step in the initiation of implantation. A receptor-­mediated paradigm for embryo attachment and invasion has long been postulated.317 Numerous endometrial and trophoblast cell-­adhesion molecules and associated moieties have been suggested as candidates to serve as attachment receptors.318–322 Historically, surface modification of the glycocalyx was a subject of much interest,323–325 but more recently, the molecules that populate the luminal surface have come under scrutiny. The actual number of receptors and ligands on this surface appears to be somewhat restricted compared with the other cells in the endometrium. As shown in Fig. 10.17, a limited number of adhesion/attachment components have been suggested. MUC-­1 (not shown) is a large glycoprotein that is downregulated on the endometrial surface in most species at the time of implantation,326 but expressed throughout the menstrual cycle in humans.322 Nevertheless, debate continues regarding this large glycoprotein as a possible attachment receptor for the human embryo. Other smaller molecules are present and may serve the purpose of initial attachment, including trophinin, integrins, CD44, L-­selectins, and HB-­EGF.327,328 The cascade of events leading to successful implantation likely requires many critical proteins with different functional

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CD44 Trophoblast

OPN 3 HB-EGF™ ErB-4 Lewis-x

Endometrium L-selectin Trophinin

contributions. A novel cell surface protein called trophinin has been suggested as a homologous pairing partner between trophoblast and endometrium during implantation.319 The ανβ3 integrin and its ligand OPN are expressed at the time of implantation on the luminal surface of receptive endometrium. Secreted OPN binds to this integrin through an RGD sequence.273,329 Since OPN can also bind to the CD44 hyaluronate receptor through non-­RGD binding sites, it has been suggested that these serve as a “sandwich”-­pairing mechanism at the point of interface.330,331 In the human, surface epithelial cells on both the embryo and endometrium express the ανβ3 integrin and CD44.332 There is also ample evidence that RGD binding is vital to the process of implantation,333,334 and peptides containing this sequence reduce implantation efficiency in animal models, including the rabbit and mouse.131,132 Thus integrin-­ mediated adhesion may somehow play a role in successful implantation. More recently, it has been suggested that OPN, ανβ3 integrin, and CD44 binding serves to suppress the innate immune system through decay-­accelerating factor (DAF)-­mediated interference with complement subunit C3,271,275 a role that could be critical for the protection of the embryo at the time of initial attachment and invasion. The transmembrane form of HB-­ EGF and its receptor, ErbB-­4, are expressed on the surface of the endometrium and on the outer cells of the embryo, respectively. Evidence in both rodents and humans suggests that these molecules could serve as an attachment receptor-­ligand pair during implantation.320,335 Soluble HB-­EGF interferes with this process, perhaps through competitive inhibition. Transmembrane HB-­ EGF could also play a paracrine role, especially if cleaved from its transmembrane location when the embryo enters the uterine cavity. Perhaps the most promising mechanism involves that previously elucidated for leukocyte/endothelial interaction. L-­selectin and an oligosaccharide ligand are expressed on the blastocyst and endometrial surface in the human, respectively.321 This type of adhesion reaction between embryo and endometrium at the time of implantation is quite appealing and likely involves integrin mechanisms for embryo invasion, similar to leukocyte intercalation at sites of inflammation. Evidence from humans illustrates a temporal and spatial regulation of L-­selectin ligand expression on the luminal and glandular epithelium.336 Likewise, each mechanism could be disrupted in infertile women, leading to a failure of implantation.337 Methods to diagnose and correct such defects may provide new hope for those with otherwise UI or RPL. Structural alterations accompany the biochemical changes noted on the surface epithelium. Scanning electron microscopy reveals that the human endometrial epithelium consists of secre tory and ciliated cells (

Fig. 10.17 Luminal surface proteins that may interact with complementary proteins on the embryonic surface.  Experimental evidence exists for each of these possible attachment reactions. It is likely that there is redundancy in these systems and that each may serve specific functions, such as signaling or attachment during the initial stages of implantation.

ciliated cells changes during the menstrual cycle, decreasing in the late proliferative phase and increasing in the secretory phase. In general, estradiol levels correlate directly with the presence of ciliated cells, and withdrawal of estrogen is associated with loss of cilia. The ciliated cells do not undergo surface morphology changes during the menstrual cycle, whereas the secretory cells display significant cycle-­dependent surface modifications. Transient surface specializations of the secretory cells called pinopodes, also known as pinopods or uterodomes, have been a focus of research because the temporal patterns of expression seem to coincide with the time of maximal uterine receptivity (see Fig. 10.18B).338 These surface structures were first identified on the luminal epithelium of rodent endometrium during the limited period (∼12 hours) when the uterus is receptive to implantation. They were shown to be involved in pinocytosis, hence the appellation of pinopode (from the Greek “drinking foot”).339 Similar structures, although with different morphologies, were subsequently discovered in numerous species, including humans; their appearance again generally correlated with the time of implantation. While it is certain that pinopodes are involved in pinocytosis in the rodent uterus, in vitro studies failed to show that they serve this function in women—hence the suggestion that the structures should be designated “uterodomes” as opposed to “pinopodes.”340,341 Mechanisms underlying the formation of pinopodes in the human endometrium have not been elucidated. They may form from the accumulation of membrane components, as a consequence of secretory activity, or from reorganization of the cell cytoskeleton. Some researchers have suggested that they serve to elevate the endometrium above the ciliated cells, providing a platform with the necessary complement of surface adhesion receptors. The role of pinopodes in human embryo implantation (beyond the temporal correlation between their appearance and the estimated time of nidation) is supported by in vitro studies, demonstrating that human blastocysts implant on human endometrial epithelial cells only in areas bearing pinopodes (Fig. 10.19). Other studies have demonstrated that surface biomarkers are present on the pinopodes. HB-­EGF, a molecule implicated as a membrane-­bound ligand and juxtacrine/paracrine factor important in signaling to the embryo, is on the surface of pinopodes at the expected time of implantation.342 The ανβ3 integrin and its ligand are also both present on these apical protrusions during the window of implantation (see Fig. 10.18D).343 MUC-­1 and OPN appear to be on different cell types of the luminal surface, based on electron immunohistochemistry; MUC-­ 1 is present solely on the ciliated cells, while OPN is present on the secretory or pinopode-­bearing cells.343 The formation of pinopodes appears to be dependent on progesterone, while estrogen causes them to regress. The

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D

Fig. 10.18 Human luminal endometrial structure. (A) The surface epithelium of human endometrium is composed of ciliated and secretory cell types. (B) During the time of maximal receptivity, the surface of these cells transforms into a sac-­like protrusion seen in many species, now called pinopodes. During the prereceptive phase, surface receptors such as the ανβ3 integrin are not present (C) but are expressed during the window of implantation on these pinopode structures (D). (Used with permission of Human Reproduction.)

earlier appearance of pinopodes in controlled ovarian stimulation cycles is correlated with the preovulatory rise in plasma progesterone. Administration of a low dose of the PR antagonist mifepristone on days 14 and 15 of a cycle delays pinopode formation. Studies in the mouse suggest that Hoxa10, a progesterone-­regulated gene, is required for pinopode formation,344 though pinopodes have been described in both Lif and Hoxa10 null mice.345 The actual temporal distribution of pinopodes has also been brought into question in the past several years, with several studies showing little correlation with the actual window of implantation.346–348 While likely involved in embryo-­endometrial interactions, the utility of these structures as markers of uterine receptivity appears limited.

Growth Factors and Cytokines Various growth factors have been implicated in the dramatic morphologic changes that occur in the endometrium during the menstrual cycle and pregnancy. Among the growth factors whose expression has been demonstrated in the human endometrium and decidua are EGF and EGF-­like molecules, including TGFA and HB-­EGF; acidic and basic FGF; IGF-­1 and IGF-­2; IL-­1, IL-­11, and IL-­6; LIF; the CSF family (CSF-­1, CSF-­2, and CSF-­3); members of the TGF-­beta superfamily; platelet-­derived growth factor (PDGF); tumor necrosis factor-­alpha (TNFA); and endothelins 1, 2, and 3. Many of these factors have been proposed to play crucial roles in endometrial function and during pregnancy.261,286,349 The endometrium and decidua and embryo/ trophoblast have also been shown to express receptors for many of these factors, including the EGF/TGFA, IGF-­1 and IGF-­2, IL-­1, CSF-­1, CSF-­2, CSF-­3, PDGF, and VEGF. With such a wide array of growth factors, it has become difficult to determine with clarity the role of each factor in endometrial growth and differentiation, or their importance in processes involving maternal embryonic interaction and placental development.

Fig. 10.19 In vitro culture of human embryos on cultured endometrial epithelium appears to show that embryos prefer to attach to areas with pinopode expression. (Reproduced with permission from Bentin-­Ley U. Relevance of endometrial pinopodes for human blastocyst implantation. Hum Reprod. 2000;15[Suppl 6]:67–73.)

Several growth factors have been shown to exert regulatory effects on the expression of extracellular matrix proteins, their cellular receptors (integrins, selectins, cadherins), and enzymes (MMPs) that also influence cell growth, differentiation, and remodeling.351–353

Leukemia Inhibitor Factor (LIF) and IL-­11 IL-6, LIF, and IL-11 are glycoproteins that belong to the same family of cytokines whose receptors utilize gp130 as a common

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signaling molecule.354,355,356 LIF acquired its name due to its capacity to inhibit the proliferation of a mouse leukemic cell line. LIF is expressed constitutively in the ampullary region of the fallopian tube and in a cyclic fashion in both epithelial and stromal cells of the endometrium, with epithelial expression being greater.357–364 The functional LIF receptor, a complex consisting of LIF receptor beta (which binds LIF) and gp130 (which mediates signal transduction) is present throughout the menstrual cycle in the luminal epithelium. LIF receptors are expressed by all trophoblast types, particularly the villous syncytiotrophoblast and cytotrophoblasts and to a lesser extent the extravillous trophoblast cells. Sentinel observations in the mouse have clearly demonstrated that LIF of endometrial gland origin is crucial for the process of implantation, particularly the decidualization response.359,365,366 LIF-­deficient female mice failed to become pregnant or respond to a uterine decidualizing stimulus. However, transfer of their embryos to pseudopregnant, wild-­type females resulted in viable pregnancies, as did infusion of LIF into the uteri of LIF-­deficient females. The primary action of LIF appears to be on the uterus. However, in mice, subsequent placentation is disrupted, perhaps because the role of LIF in modulating trophoblast differentiation and expression of MMPs cannot be executed. Implantation of LIFR-­deficient human embryos occurs but these offspring suffer from Stuve-­Wiedemann/Schwartz-­Jampel type 2 syndrome.362 LIF mRNA and protein are present in human endometrium, being most abundant in the glandular and luminal epithelium and peaking in the secretory phase of the cycle.367,368,369 The cycle-­ dependent expression of LIF in human endometrium may be a function of other growth factors, rather than a direct effect of steroid hormones. If implantation occurs, LIF expression by the endometrial glands is downregulated, while there is a concomitant increase in LIF expression by endometrial NK and T cells. LIF has been found to enhance human blastocyst formation and modulate trophoblast differentiation in vitro.358 LIF levels in uterine flushings rise 7 days after ovulation, reaching a maximum 5 days later. LIF levels in uterine flushings and secretion of LIF from cultured endometrium obtained from patients with repeated implantation failures or UI is decreased357,370 and defects in LIF have been implicated in some cases of RPL.358 Leptin has been suggested as a regulator or LIF, through its receptor, OB-­R.371 LIF stimulates hCG in trophoblast.363 LIF has been implicated in activation of STAT-­3; mice homozygous for STAT activation site on gp130 have a defect nearly identical to LIF deficient mice.372 Suppressor of cytokine signaling protein-­3 (SOCS-­3) is stimulated by LIF and blocks phosphorylation of gp130 and STATs. In LIF null mice, stromal PTGS2 and epithelial HB-­EGF are absent at the site of implantation.373,374 Two other EGF family members, AMP and epiregulin (EPR), are also reduced in LIF knockout mice. IL-­1 has been shown to induce PTGS2.375,376 Prostacyclin acting through the PPARγ is essential for decidualization.377,378 Cox-­2 (or Ptgs2) null mice have multiple defects in ovulation, fertilization, and implantation.379 Together, LIF stimulation of luminal epithelium along with blastocyst may trigger IL-­1 that triggers decidual changes. Mutations in the coding sequence of one copy of the LIF gene were identified in a small number of infertile women (3 of 74 infertile nulligravid subjects), and a presumed polymorphism was found in one of 75 fertile controls and none of 131 nonobstetrical patients.380 One of the mutations in the infertile group was in the 5′-­regulatory region of the LIF gene; the two others were in the coding sequence in a domain thought to be important for LIF binding to its receptor. Unfortunately, whether LIF levels or bioactivity in uterine flushings or endometrial biopsy material were correlated with genotype was not determined. Collectively, however, these observations are consistent with an important role for LIF in human implantation, trophoblast differentiation, or placentation. IL-­11 is another member of the IL-6 family that is implicated in the decidual response.

knockout mice, female mice lacking the IL-­11 receptor α-­chain are infertile because of defective decidualization.381 IL-­11 is present in the human endometrium, and it advances progesterone-­ induced decidualization of cultured endometrial stromal cells. Both relaxin and prostaglandin E2 increase IL-­11 expression. An inhibitor of IL-­11 (W147AIL-­11) reduces prolactin secretion by ESC in response to RLX and PGE2, suggesting IL-­11 is the critical factor in this signaling cascade.383 Like LIF, IL-­11 is an activator of the JAK/STAT signaling pathway through STAT3, which stimulates the suppressor of cytokine signaling-­3 (SOCS3), a negative feedback mechanism for receptor activity. Ovarian steroids and cAMP differentially stimulate STAT3 and SOCS3, respectively, in vitro, while IL-­11 activates both by phosphorylation.384 Treatment with an antiprogestin increases SOCS3, attenuating IL-­11-­induced STAT3, suggesting multiple regulatory components, including progesterone. Recent investigations in women suggest that IL-­11 and phosphorylated STAT3 are significantly lower in the infertile endometrium compared with controls, while IL-­11 receptor and LIF were not different.385

Epidermal Growth Factor Family of Growth Factors The EGF family of growth factors appears to play a major role in uterine development and physiology.261,286 The EGF family of ligands is produced by a family of genes, EGF, HB-­EGF, AMP, betacellulin (BTC), EPR, TGFB, and neuregulins (NRG). This EGF family of ligands has the ability to bind and activate one or more of four homologous ErbB receptors via a conserved 60 amino acid “EGF-­like” binding domain. These four receptors differ in their activities and dimerize with each other to further refine the diversity of this growth factor family. Interacting with their receptors, the EGF family acts as autocrine and juxtacrine factors, and some exist as membrane-­bound forms that are released by proteolytic cleavage to function in a paracrine or endocrine manner. The proteolytic cleavage is accomplished by cell-­surface metalloproteinases, similar to those involved in L-­selectin ligand and MUC1 cleavage, suggesting an important role for metalloproteinase action at the maternal-­fetal interface.386–388 The receptors for the EGF ligand family, ErbB1, ErbB2, ErbB3, and ErbB4, are structurally homologous, and the specificity of ligand binding is determined by differences in extracellular domain sequences (Fig. 10.20). Molecular studies have determined that these receptors are regulated through the IHH pathway, discussed earlier.389 The ErbB proteins function as hetero-­or homodimers and possibly also as higher-­order multimers.390 ErbB2 lacks ligand-­binding activity, and ErbB3 lacks functional tyrosine kinase activity. Thus ErbB2 and ErbB3 likely function only as heterodimers, with another ErbB protein supplying the missing function in a dimer. EGF and TGFA stimulate the proliferation of endometrial stromal cells.261 The synthesis of fibronectin and vitronectin by epithelial cells is enhanced by EGF. EGF also stimulates stromal cell differentiation and enhances the synthesis of laminin and fibronectin. These growth factors also enhance the morphologic and functional differentiation of decidual cells in vitro. EGF has not been identified in the circulation of cycling or menopausal women, but it has been detected in the serum of pregnant women, with peak concentrations occurring in early pregnancy. Binding studies indicate that EGFR peaks at the time of ovulation (or shortly thereafter) and declines during the secretory phase to a nadir immediately before menses. The binding sites are present on both stromal and epithelial cells, as well as in the decidua of pregnancy. Studies using immunohistochemistry suggest that decidualization is associated with an increase in EGFR. Abnormal EGF and EGFR activity has been reported in cases of intrauterine growth retardation. Since IHH regulates the EGF receptors, progesterone resistance associated with inflammatory conditions such as endometriosis could account for defects in

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract EGF AMP TGF

BTC HB-EGF EPR

NRG1

NRG1-4

Plasma membrane

Tyrosine kinase

X

ErbB1

ErbB2

ErbB3

ErbB4

Fig. 10.20 The epidermal growth factor (EGF) family of growth factors and receptors.  EGF and its family members associate with specific receptors that signal through intracellular tyrosine kinase action. The four receptors can dimerize in predictable ways.

stromal proliferation and decidua formation noted in this condition.389 It has been hypothesized that EGF and related molecules play a role in the induction of trophoblast differentiation. Multiple EGF family members appear to directly participate in the process of implantation.391 HB-­EGF exists as a membrane-­ anchored precursor (HB-­EGF) that gives rise to a soluble processed form. These proteins can bind to two different EGF receptors, HER1/ErbB1 and HER4/ErbB4. HB-­EGF mRNA is found in the stroma during the proliferative phase, but in the midsecretory phase, it is also detected in the luminal and glandular epithelium.392,393 This expression pattern appears to be driven by a combination of progesterone and estrogen. HB-­EGF is postulated to play roles both in adhesion via the membrane-­ anchored precursor (which binds to HER4/ErbB4 on the apical surface of the trophectoderm) and in stimulating embryo growth. In the rodent, HB-­EGF is one of the first cytokines found around the implanting embryo.335,394 In the human endometrium, mRNA levels of HB-­EGF increase during the secretory phase, reaching a peak at the time of implantation.392 HB-­EGF can mediate the attachment of human embryos in an in vitro assay of implantation.320 HB-­EGF also may promote embryonic development. When added to a serum-­free medium, it increases the number of embryos reaching the blastocyst stage and stimulates hatching. HB-­EGF may also participate in paracrine actions within the endometrium. Stromal-­derived HB-­EGF promotes expression of LIF, HOXA10, ανβ3 integrin, and DAF by endometrial cells in vitro. Other members of the EGF family likely play important roles during early pregnancy as well.357

Transforming Growth Factor Beta Family The transforming growth factor beta (TGFB) family, multifunctional proteins that regulate cell growth and differentiation, includes five dimeric polypeptides encoded by related genes.278,349,395,396,397 These growth factor members bind to three cell-­surface proteins designated type I, II, and III receptors. The type I and type II receptors are thought to mediate the actions of TGFB through the SMAD signaling pathway. All three isoforms of TGFB can be found in human endometrium, including TGFB1, TGFB2, and TGFB3. Null mutant mice bearing TGFB mutations for each of the three genes produce a distinct phenotype suggesting

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a separate role for each member of this growth factor family.398 All three and the type II receptors were identified in all cell types of the endometrium.399 TGFB1 and TGFB3 are found on both epithelial and stromal cells, while TGFB2 is found primarily on the stroma and increases during the secretory phase.400–402 TGFB3 increases at menstruation and remains high throughout the proliferative phase, while TGFB1 is maximal at menstruation. Ovarian steroids strongly repress TGFB2 and TGFB3 in the stroma, but only TGFB2 is repressed in the glandular epithelium. cAMP prevents this inhibition of TGFB2 by progesterone, while MAP kinase (MAPK) inhibitors differentially stimulated TGFB2 and TGFB3. These opposite effects provide insight into the discreet and highly temporal and spatial expression of the TGF cytokines during the menstrual cycle. TGFB has profound effects on the extracellular matrix, increasing the synthesis of collagen while decreasing its degradation.158,403 In the secretory phase, TGFB is a potent inhibitor of MMP activity in both stroma (MMP-­3) and epithelium (MMP-­ 7), while simultaneously promoting TIMP expression. This cytokine is induced by progesterone, acting in concert with retinoic acid (RA) to maintain these profibrogenic activities. In the event of pregnancy, progesterone acting through TGFB will maintain the integrity of the endometrium throughout gestation. In lieu of pregnancy, the fall in progesterone releases the MMP inhibitory factors, resulting in a programmatic breakdown of the extracellular matrix and subsequent menstruation. Other members of the TGFB family are expressed in the endometrium.349 EBAF (or LEFTY) may participate in the menstruation process, acting as a potent inhibitor of TGFB.221 Antagonism of TGFB1 by overexpression of EBAF (Lefty; a TGFB antagonist) reduces the number of implantation sites in rodent models,404 consistent with human data showing high levels of LEFTY in infertile women.224 Activins are highly abundant in the endometrium. Activins beta (BA and BB) is found in the glandular epithelium with peak expression during the secretory phase. Activin A promotes decidualization in vivo, while follistatin opposes the actions of activin.405 Activin A enhances the production of LIF and enhances MMP2 during decidualization. MMP and LIF activity are both crucial for decidualization in rats and primates. TGFB1 and TGFB2 have been reported at the interface between embryo and endometrium and likely play multiple roles during embryo attachment and invasion.397,406 In pregnancy, TGFB is most abundant in the decidua in the first trimester of pregnancy, where it is thought to restrain trophoblast invasion by promoting trophoblast differentiation away from the invasive phenotype. TGFB is sequestered in the extracellular matrix, where it can be activated by embryo-­derived proteases. TGFB upregulates the expression of cellular fibronectin by trophoblast cells and induces integrins on the trophoblast that interact with the ECM during an invasion, promoting MMP activity.407,408 TGFB induces TIMPs and PAI-­1 activity to counterbalance the invasiveness of the trophoblast, and TGFB is a potent immunosuppressant that may prevent the maternal immune rejection of the fetal allograft. Its actions include suppression of chemotaxis and macrophage and T-­cell activity. ADAMTS (a disintegrin and metalloproteinase with thrombospondin repeats) has been thought to play a critical role in extracellular matrix (ECM) remodeling during implantation. IL-­1 increases, while the antiinflammatory effects of TGF-­β1 decrease, ADAMTS expression.409 These opposing actions illustrate the balance between factors favoring embryonic invasion and maternal efforts to control this invasion and reflect the essential role of TGF-­β in this process.

Other Growth Factors PDGF is produced by endometrial stromal cells and also released PDGF is a potent

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mitogen that acts on endometrial receptors that are most abundant in the proliferative phase of the cycle. FGFs encompass a family of growth factors that can stimulate growth of endometrial cells and smooth muscle cells.411,412 Acidic and basic FGFs bind to proteoglycans. Because these proteins do not contain secretory signal sequences, they may be most important in the endometrium during menstruation when they could be released from dying cells. Basic FGF is angiogenic but also stimulates stromal cell proliferation in the presence of progesterone. FGF-­7, also known as keratinocyte growth factor, stimulates epithelial cell proliferation. FGF-­7 mRNA is expressed at the highest level in the late secretory phase endometrium in the stroma; its receptor is most abundant in the glandular epithelium in the late proliferative phase. These findings suggest that FGF-­7 is progesterone dependent, whereas its receptor is estrogen sensitive. FGF-­8 is a uterine growth factor isolated from a bovine uterus, and FGF-­9 was recently described in the late proliferative phase of human endometrium.412 The IGF system of growth factors encompasses not only IGF-­1 and IGF-­2 hormones but also two distinct receptors and multiple different binding proteins that modulate IGF activity.262,263,413–417 IGF-­1 and IGF-­2 are mitogenic growth factors that share structural similarities to insulin and are present in the mouse uterus and throughout the menstrual cycle in women.418 Both hormones are present in the stroma, though the receptors are present in both epithelial and stromal cells. IGF-­1 is more abundant in the proliferative phase and may function in epithelial proliferation, while IGF-­2 is expressed in a more robust fashion in the secretory phase and is thought to be a mitogen for the stroma of pregnancy. IGF-­1 binds to both the type 1 IGF receptor (IGF-­1 receptor), structurally similar to the insulin receptor and the type 2 receptor (IGF-­2 receptor) with high affinity. IGF-­2 binds the latter with higher affinity.419 IGF-­1 mRNA is localized by in situ hybridization primarily in the syncytial trophoblast. IGF-­2 mRNA has been demonstrated in mesenchymal fibroblasts of the villous core and is also expressed in first-­trimester and term cytotrophoblasts. The expression of IGFs in the placenta seems to be under the regulatory control of hormones (insulin, human placental lactogen, and estrogens) as well as growth factors, including PDGF. Type IGF-­1 receptors have been detected in the placenta during the earliest periods of gestation, and it has been hypothesized that IGFs promote trophoblast proliferation. TNFA is a pleiotropic factor that exerts inflammatory, mitogenic, mitostatic, angiogenic, and immunomodulatory effects in a variety of tissues.420 It is a membrane-­bound 14-­kDa polypeptide that is derived by proteolytic cleavage from the 26-­kDa precursor. Expression of TNFA mRNA and protein has been demonstrated in human endometrium, decidua, and trophoblasts; its receptors have also been found in all these tissues. TNFA in the human endometrium is subject to regulation by steroid hormones—namely, estrogens and progesterone. TNFA and its receptors (TNF-­R) are expressed by trophoblasts during both early and late gestation.421 There is differential expression of the two genes encoding TNF-­R, thus allowing some regulation of TNFA activity. In cultured human chorionic cells, TNFA affects cellular fibronectin secretion. Scientists have hypothesized that, along with other endometrial and trophoblast factors, TNFA controls trophoblast adhesion and invasion and alters the integrin expression pattern in endometrial stroma.422,423 Colony-­stimulating factors are a family of three cytokines, CSF-­1, CSF-­2, and CSF-­3, that were initially identified by their ability to stimulate hematopoietic stem cells to form colonies in semisolid culture media, with CSF-­1, 2, and 3 promoting proliferation and differentiation of macrophage, granulocytes plus macrophage, and granulocytes, respectively. The original observation that short-­term cultures of human placental tissue pro duced large amounts of CSFs

factors in trophoblast and, more recently, in the endometrium. Despite growing enthusiasm for clinical therapy with CSFs to improve implantation, human data on their normal endometrial expression and function are minimal. CSF-­ 1 gene expression is essential for normal fertility in mice.425,426 Expression of CSF-­1 and its receptor are found in the endometrium, decidua, and placenta, with a higher CSF-­1 expression in the secretory phase and decidua than in the proliferative phase.427 The CSF-­1 receptor is highly expressed by extravillous trophoblasts in the cell columns that anchor the placenta to the uterus, with nearby decidual cells expressing CSF-­1, consistent with in vitro data demonstrating CSF-­1 effects on trophoblast proliferation and differentiation.428 The phenotype of the op/op (osteopetrotic) mouse that lacks CSF-­1 argues against an essential role for this factor, as the observed reduced fertility and smaller litter sizes are due, in large part, to an ovulation defect, though placental abnormalities are evident in a CSF-­1 knockout.425,426 Thus, despite the production of CSF-­1 by the endometrium, the presence of CSF-­1 receptor on the trophoblast, and clear evidence of CSF-­1 effects on trophoblast function, a definitive role for endometrial CSF-­1 remains to be established. CSF2 and its receptor are also produced by both the placenta and endometrium. CSF-­2 has been shown to increase the proliferation of trophoblast cells in vitro and murine Csf2 null placentae demonstrate altered morphology and gene expression.429 As is the case for CSF-­1, a significant role for CSF-­2 in human reproduction remains unclear. CSF3 is a monomeric glycoprotein, originally characterized as a factor promoting the production, differentiation, and function of granulocytes. Although CSF3 and its receptor are also produced by the endometrium and placenta, the details of this expression are not well described. Despite the lack of information, CSF3 has been shown to be an effective treatment in recurrent miscarriage and implantation failure, each in a single randomized trial.430,431 Although confirmatory studies are needed, initial data suggest that targeting the maternal CSF3 receptor may be an important therapy for some infertile couples.

Human Chorionic Gonadotropin In vitro studies demonstrate that the addition of follicle-­ stimulating hormone, luteinizing hormone (LH), human chorionic gonadotropin (hCG), thyroid-­stimulating hormone, and free beta subunit has an effect on human reproductive tract tissues, including the stimulation of prolactin production, enhancement of decidualization, and myometrial relaxation.432,433 The presence of LH/hCG receptors in the fallopian tube, myometrium, and endometrium has also been reported. However, the transcripts detected are smaller than those encoding the gonadal LH/hCG receptors; the proteins are also smaller, detected as 50-­to 60-­kDa molecules as compared with the observed receptor molecular weights in gonadal tissue (83 to 95 kDa). Thus the extragonadal LH/hCG receptors appear to be truncated, evidently lacking extracellular domains but still retaining the capacity to signal after binding LH and hCG. The primary signal transduction cascade initiated by the truncated endometrial LH/hCG receptors may not involve the classical cAMP/protein kinase A system as the primary pathway but rather the mitogen-­ activated protein kinase pathway or the activation of prostaglandin synthesis. Although extragonadal LH/hCG receptors have been proposed to play various roles in the reproductive tract, they are probably most important in the context of pregnancy, where high levels of hCG are present. Thus hCG could be an important embryonic signal in the bidirectional dialogue between the conceptus and the uterus. The best evidence that extragonadal LH/ hCG receptors have an important role in primate reproduction is derived from the study of the baboon, where the administration

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract

of recombinant hCG causes alterations in both epithelial and stromal cells. Preliminary studies in women indicate that hCG can also affect the human endometrium in vivo,434 including prevention of endometrial apoptosis.434,435 In clinical trials, hCG support has been shown to decrease miscarriage, perhaps by this mechanism of CL rescue and avoiding loss of CL support to the nascent pregnancy.436 In the baboon, the response to intrauterine administration of hCG by osmotic minipump over a 4-­day period starting on day 6 after ovulation includes the formation of an epithelial plaque, hypertrophy of the surface epithelium, and rounding off of the glands characteristic of pregnancy.433 Glycodelin expression in the endometrial glandular epithelium is enhanced, and alpha smooth muscle cell actin is expressed in the stroma. In vitro, hCG inhibits stromal cell apoptosis and stimulates decidual changes, reflected by increased IGFBP-­1 expression as well as increased expression of PTGS2. As previously noted, in vitro studies have shown that the glycoprotein α subunit stimulates the production of prolactin, another uterine secretory product. Activation of myometrial LH/hCG receptors in vitro causes myometrial relaxation, which could facilitate nidation in vivo.434 The response to hCG was blunted in animals with endometriosis .437 In vivo studies have demonstrated a direct effect of hCG on endometrial histology and receptivity.438,439 Using intrauterine microdialysis in the human, it was shown that administration of hCG significantly reduces IGFBP1 and M-­CSF expression after postovulatory day 10, while LIF, VEGF, and MMP9 were all substantially increased,268,440 suggesting that hCG is an important modulating factor during early pregnancy. In vitro studies, using hCG coated beads, have demonstrated an induction of trophinin on the endometrial epithelium when IL-­1 is present, suggesting a mechanism for enhanced embryo/endometrial interactions at the maternal-­fetal interface.441 LIF and its receptor are upregulated in response to hCG treatment in vitro.442 As an endocrine factor, hCG likely has independent effects on a wide variety of endometrial genes, independent of direct trophoblast interactions.269

Prostanoids and Other Lipids The role of prostaglandins in the implantation process has long been suspected because of their effects on the vascular system and their association with inflammatory processes (see Chapter 4). The fact that mice deficient in Ptgs2 display abnormalities in the implantation process—particularly the early decidual response—is consistent with this notion.434 Evidence suggestive of a role of prostanoids in human implantation includes the presence of PTGS1 and PTGS2 in the human endometrium (mainly the glandular epithelium) during the presumptive implantation period;443 an examination of prenatal use of nonsteroidal antiinflammatory drugs (NSAIDs), which inhibit COX enzymes, indicated an increased risk of miscarriage in users of aspirin and other NSAIDs.444 One of the key prostanoids involved in implantation is thought to be prostacyclin, a ligand for the peroxisome proliferator-­activated receptor delta (PPARD), a nuclear receptor family member expressed in subluminal stromal cells in the rodent uterus.377,445–447 This transcription factor is implicated in the implantation process as well. Another lipid implicated in implantation is the arachidonate derivative known as arachidonoylethanolamine or anandamide, a ligand for the cannabinoid receptors.448–451 Anandamide, referred to as an endocannabinoid, binds to cannabinoid receptors CB1-­R and CB2-­R, which are expressed in the preimplantation embryo and in the reproductive tract. The embryo is enriched in CB1-­ R, and in the blastocyst, the expression of this receptor is most abundant in the trophectoderm. Uterine anandamide levels in the mouse are reduced at the time of implantation and are highest at interimplantation sites. Endocannabinoids at low levels accelerate trophoblast differentiation, but at high levels inhibit

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trophoblast differentiation and arrest embryo development. Tetrahydrocannabinol and synthetic agonists of the cannabinoid receptors have similar effects. Thus it has been postulated that endocannabinoids play an important role in controlling the synchrony of embryo development for implantation in rodents. Anandamide is present in human reproductive tract fluids,450 and high anandamide levels are associated with in vitro fertilization (IVF) failures.451

Immunology of the Endometrium The uterus is an immunologically privileged organ: it can accommodate tissue invasion by immunologically semiforeign placental cells, yet maintain mucosal immune defenses against ascending foreign organisms, and provide a system to efficiently clear the endometrial detritus that results from menstruation (see Chapter 13). Remarkably, the endometrium also uses mechanisms of acute inflammation during normal, hormonally regulated physiologic processes, including menstruation and embryo implantation. These acute inflammatory episodes are quickly resolved, avoiding the consequence of scarring and dysfunction. Interestingly, the induction of acute inflammation through the endometrial scratch technique has been adopted into clinical practice in an attempt to increase pregnancy rates following IVF treatment; however, there is a lack of randomized clinical trials to support any clinical benefit.452,453 Despite the description of critical active processes to resolve inflammation in other tissues and the clear relevance to endometrial physiology and pathophysiology, mechanisms that resolve endometrial inflammation and their role in pregnancy establishment remain largely unstudied.454,455 The complex requirements of uterine immunity and tolerance use overlapping and redundant mechanisms dependent on both innate and adaptive branches of the immune system. Endometrial immune processes, as with other uterine functions, but unlike those for other mucosal immune sites, are markedly influenced by cyclic and pregnancy-­specific changes in sex steroid concentrations and possibly by hCG.456 The endometrium is populated by BM-­derived immune cells, as well as endometrial epithelial and stromal cells that regulate immune function. As is the case for many epithelial cells, endometrial epithelium expresses members of the Toll-­like receptor family (TLR2 to 6, 9, and 10), which detect pathogen products and trigger a cellular response to these “foreign” molecules, including peptidoglycans from Gram-­positive bacteria (TLR2), lipopolysaccharide from Gram-­ negative bacteria (TLR4), and unmethylated CpG islands found in bacterial DNA (TLR9). The endometrium also produces host defense molecules, defensins, as well as cytokines and chemokines. Uterine lymphoid and myeloid cells play roles in tissue defense, immune modulation, angiogenesis, and tissue remodeling.153,457–460 These cells are present in the fallopian tubes, uterus, and cervix, with the fallopian tubes and uterus containing a higher proportion of leukocytes than the cervix and vagina.461 The endometrial innate and adaptive immune systems are regulated by steroid hormones. For example, progesterone induces a local Th2-­type cytokine response in the uterus, which includes an increase in IL-­4, IL-­5, and IL-­15 and downregulation of the IL-­13 receptor alpha 2, which is a negative regulator of the antiinflammatory cytokine, IL-­13, and powerful inhibitor of the Th2 response.462 The Th2 response is believed to counter proinflammatory processes in the endometrium that could lead to rejection of the embryo. Steroid hormone-­directed alterations in endometrial chemokine production influence the trafficking of blood leukocytes in the reproductive tract. Furthermore, the actions of progesterone are important in the overall immune suppressive phenotype adopted by the receptive endometrium.463 During the secretory phase, there is a profound recruitment of leukocytes into the endometrium starting in the perivascular

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locations around spiral arterioles and glandular epithelium.464–467 The progesterone-­ induced alteration in endometrial cytokine/chemokine production contributes to this recruitment.468 Cytokines IL-­ 1, IL-­ 11, IL-­ 15, LIF, and TGFB regulate the trafficking of leukocytes to the endometrium.469 IL-­15 recruits NK cells into the endometrium,470 and IL-­15 knockout mice lack uNK cells. Locally acting prostaglandins (PG), including PGE along with VEGF, modulate vascular permeability.467,471 Cyclooxygenase-­2 (COX-­2), a rate-­limiting enzyme that regulates the biosynthesis of PGE2, is critical to implantation in the mouse,378,379 and in animal models, PG are required for initiation and maintenance of decidualization.472,473 Blockade of COX-­2 prevents decidualization in mice379 and clearly plays a role in endometrial function surrounding pregnancy474,475 with reduced COX-­2 associated with implantation failure476 and RPL.477 An immunotolerance maternal immune response is essential for the acquisition of endometrial receptivity and the success of pregnancy.478–480 Factors that support a more suppressed immune environment, including the recruitment of T regulatory cells (Tregs) and shunting away from a proinflammatory, Th1/ Th17 responses are central to our understanding of infertility and pregnancy loss associated with various inflammatory conditions such as endometriosis, as discussed later.

regulating complex and coordinated biochemical interactions in the endometrium that supervise the controlled invasion of the fetal allograft, including direct involvement in the establishment of immunologic tolerance.286,483,484 The actions of P are mediated in part through RA and TGFB485–487 and PPARG, with a collective profound influence on CD4+ leukocyte differentiation. The actions of progesterone (and RA and PPARG) are disrupted in certain gynecologic disorders, including endometriosis, leading to alterations in the endometrial phenotype and contributing to infertility and early pregnancy loss.328,488 Normal eutopic endometrium hosts leukocytes in both stromal and intraepithelial locations464,466 and an immunotolerant environment appears essential for successful implantation. Recent evidence suggests that maternal immune responses are critical for the acquisition of endometrial receptivity.478–480 and that immune cells can directly regulate endometrial epithelial receptivity and associated gene expression.117,489 Macrophages have been shown to regulate uterine epithelial gene expression in the mouse,490 suggesting a mechanism for observed differences in eutopic endometrial gene expression differences, including progesterone resistance seen in endometriosis. Conversely, inflammatory cytokines in peritoneal fluid of endometriosis might alter the differentiation pathways of monocytes to a more hostile phenotype.491

Leukocytes and Lymphocytes

T-­Regulatory and TH-­17 Cells Regulate Endometrial Receptivity

There are striking menstrual cycle-­dependent changes in the immune cell population of the endometrium. Neutrophils, the most abundant leukocytes of the immune system, are rare in the normal endometrium until the perimenstrual phase, when they accumulate and account for 6% to 15% of the total cell number. Eosinophils are also rarely found in the normal endometrium until the perimenstrual phase, when they also accumulate—usually in aggregates—and show evidence of activation, as revealed by the extracellular location of eosinophil cationic proteins. Macrophages are present in the endometrium throughout the cycle, increasing in number from the proliferative to the menstrual phases. Immediately prior to menstruation, the number of macrophages in the functional layer of the endometrium is equivalent to that of neutrophils. Mast cells expressing tryptase (a characteristic of mucosal mast cells) and chymase (a characteristic of connective tissue mast cells) are found in the human endometrium. In the functionalis, mast cells are positive only for tryptase, while those in the basalis express both enzymes. Degranulation of these cells may initiate activation of MMPs as a result of tryptase and chymase action on proMMP-­1 and proMMP-­3. The endometrial lymphoid system has a distinctive composition and activity. There is good reason to believe that endometrial T cells are activated in situ, as judged by their expression of antigens that are characteristic of the activated state; these antigens include the MHC class II molecules HLA-­DR, HLA-­DP, and HLA-­DQ, and very late antigen 1. CD3+ T cells represent only 1% to 2% of the lymphomyeloid cells detectable in the human endometrium. They are present throughout the cycle in aggregates in the basalis, as well as singly throughout the stroma and in intraepithelial sites. The number of these cells increases prior to menstruation. The ratio of CD4+ T helper lymphocytes to CD8+ cytotoxic T cells in the endometrium is inverted compared with peripheral blood. The latter have cytolytic activity during the proliferative phase, with much reduced secretory phase activity. Few B cells and plasma cells are present in the human endometrium. Leukocytes make up between 15% and 30% of the cellular mass of the endometrium.481 The complex network of cytokines produced directly or indirectly in response to progesterone likely is involved in leukocyte recruitment to the endo metrium.469 Progesterone (P) is essential for implantation,

As discussed in Chapter 13, the immune cells can be classified as belonging to the innate branch, including monocytes, macrophages, dendritic cells (DC), neutrophils, basophils, mast cells, and lymphoid NK cells. The adaptive branch immune cells include the T and B cells, which generally require innate immune cells for activation and establishment of immunologic memory.492 BM-­derived cells are known to traffic to the endometrium via steroid-­regulated chemokine production.177,493 Under the proper circumstances, DC and Treg cells increase at the implantation site, along with uNK cells that interact with the invading placental cells to both direct and limit trophoblast invasion.494 BM-­derived decidual cells ultimately determine pregnancy outcomes.488,495–503 Monocyte-­derived leukocytes and uNK cells comprise up to 20% to 40% of all endometrial cells in the secretory phase and are critical to the establishment of endometrial receptivity.465,494,504 A loss of function may account for UI and RPL, or in women with minimal and mild endometriosis.479,484,488,505–507 Adaptation of innate immunity is required for pregnancy and survival of the semiallograft.484,488 This immunotolerance is mediated in part by T-­regulatory (Treg) cells.505,508 Treg cells are suppressors of inflammatory immune responses that keep the immune system in control.509 In the mouse model, Treg cells proliferate and are activated in the uterus in response to paternal antigens and nonspecific hormonally regulated factors.488,510 In contrast to Tregs are Th17 cells, a proinflammatory T cell subtype associated with multiple autoimmune and inflammatory conditions including colitis, multiple sclerosis, and rheumatoid arthritis.511 IL-­ 9 induces Th17 cell differentiation,512 and in mouse models of inflammatory disease, attenuation of IL-­9 decreases Th17 cells and reduces IL-­6 producing macrophages,513 contributing to Th17 differentiation. This balance in T-­cell differentiation may ultimately define whether an endometrium is receptive to the fertilized oocyte and whether a pregnancy will be successful.

Uterine Natural Killer Cells The uterine NK cell, also known as the granular lymphocyte, is a unique member of the lymphoid lineage found in the endometrium.457 These are round cells that characteristically have bilobed or indented nuclei and a pale cytoplasm containing

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract

acidophilic granules. They are a specialized subset of NK cells based on their expression of cell surface antigens (CD3−, CD16+, and NCAM/CD56bright) that is different from blood NK cells. Uterine NK cells are among the most abundant lymphomyeloid cells in the perimenstrual endometrium. Very few are present in the proliferative phase of the endometrium; they accumulate during the secretory phase, at which time they comprise 15% to 25% of the cells in the endometrial stroma. It has been postulated that the dynamic changes in uterine NK cell number during the cycle are determined by changes in endometrial prolactin production, which is then controlled by ovarian steroid hormones. This notion is based on the known roles of prolactin as an immunomodulator and the correlation between endometrial prolactin levels and NK cells.514,515 The NK cells persist in the decidua during the first trimester, when they represent 70% of the decidual leukocyte population. Their close association with stromal cells has led to the suggestion that they have an important role in initiating and maintaining decidualization. In a nonfertile cycle, the NK cells that amass during the secretory phase undergo programmed cell death. When activated IL-­2 in vitro, the NK cells become competent to kill malignant cells (and some normal ones) through the release of cytotoxic proteins like perforin. From the late proliferative phase on, the NK cells express cytotoxic activity and are thought to play a role in protecting the endometrium against infection. The cells have also been proposed to modulate trophoblast invasion during implantation and placentation because of their abundance in the decidua during the first trimester. Although in vitro studies document the killing activity of cytokine-­activated NK cells, there is little evidence for destruction of trophoblast cells in vivo. Thus any restraining activity that NK cells exert on invading trophoblast cells appears to be through a noncytotoxic mechanism that may involve secreted cytokines, including colony-­stimulating factor-­1 (CSF-­1), IL-­1, LIF, and interferon gamma (IFNG). The apparent in vitro and in vivo resistance of trophoblast cells to death may be explained by the fact that trophoblast cells express a nonclassical and nonpolymorphic major histocompatibility complex (MHC) class I antigen, HLA-­G; NK killing activity is also suppressed by endometrial stromal cells.

Innate Lymphoid Cells In addition to NK cells, other uterine innate lymphoid cells (ILCs) have been recently examined in cycling human endometrium and decidua.516,517 Three types of ILCs have been described, but these studies identified only type 3 ILCs in human endometrium. Type three ILCs seem to play a role in in-­gut immunity and mucosal homeostasis, and these cells play an important role in innate immune colitis, suggesting potential roles in endometrial inflammatory regulation.518 However, an important role for these cells in endometrial physiology or pathology remains speculative.

Regulation of Endometrial Immune Cells The changes in lymphomyeloid cell populations in the endometrium, particularly their accumulation in the premenstrual phase, appear to be the result of recruitment from the peripheral circulation and intraendometrial proliferation. The accumulation of migratory cells is directed by chemoattractant cytokines, chemokines, and the expression of intercellular adhesion molecules that attach leukocytes to the endothelium in preparation for extravasation and trafficking through the endometrium.519 Expression of these molecules changes during the menstrual cycle, at least partly under the influence of steroid hormones. The expression of several chemokines has been documented in the human endometrium including fractalkine (CX3CL1), RANTES (CCL5), IL-

245

eotaxin (CCL11). These chemokines bind to receptors on leukocytes, promoting the appearance of molecules that mediate adhesion to the endothelium, extravasation, and chemotaxis along the concentration gradient of the chemokine, thus targeting specific leukocyte types to specific endometrial compartments. A repertoire of cell adhesion molecules expressed in the endometrium, including ICAM-­1, ICAM-­2, VCAM-­1, E-­selectin, and PECAM, specifies the recruitment and location of leukocyte and platelet accumulation. ICAM-­1 is present in the functionalis in the menstrual phase; ICAM-­2 is restricted to the vascular endothelium and does not appear to change during the cycle, VCAM-­1 and E-­selectin appear in the upper layer of the functionalis in the secretory phase, and the platelet endothelial cell adhesion molecule PECAM is abundant in the stroma during the menstrual phase. Evidence for leukocyte proliferation in the endometrium includes the expression of Ki67 and incorporation of BrdU (both proliferation markers) by NK cells with a marked increase in the secretory phase. The uterine NK cell proliferation may be driven by progesterone-­regulated stromal factors as CD56+ cells proliferate in vitro in the presence of progesterone-­treated endometrial tissue. LIF is associated with a shift in Th1 to Th2 shift that is required for normal pregnancy and is defective in recurrent abortions in mice.520 Maternal T cells increased LIF expression in response to paternal antigens and trophoblast, and the reduced response was associated with poor reproductive outcomes.521 Progesterone is an antiinflammatory immunomodulator in pregnancy and through induction of LIF and other cytokines that support trophoblast invasion.463 LIF and another biomarker of endometrial receptivity, ανβ3 integrin, are coexpressed in mouse and human endometrium,342,522 and like ανβ3 integrin, disturbances in LIF expression are associated with multiple reproductive pathologies.354,357,523 While recent reports have linked these two important biomarkers,524,525 their coregulation has only recently been demonstrated.526 Absence of both integrin and LIF is associated with decreased pregnancy success in IVF.527–529 Injection of PF from women with endometriosis reduced both LIF and ανβ3 integrin expression in the mouse uterus.522 Both LIF and ανβ3 integrin are positively regulated by paracrine actions of progesterone such as HB-­EGF, and negatively regulated by inflammatory immune cells in the eutopic endometrium associated with endometriosis522,530 and inflammatory tubal disease.531 LIF has antiinflammatory actions regulating cytokine action,532 protecting against septic shock in the mouse model,533,534 whereas ανβ3 integrin has been shown to interact with decay accelerating factor (DAF) and OPN,328,535 which together block complement activation536 as part of innate immunity. T cells from women with RPL secrete less LIF than normal women, and defective LIF is associated with decreased Th2 cells and RPL.520,537

Complement System The human fallopian tube, endometrium, and cervical mucosa express components of the complement system.538–541 The activation of the third (C3) and fourth (C4) component of complement induces chemotaxis of inflammatory cells, enhances phagocytosis, and mediates cell lysis. This system of natural immunity must be tightly regulated so that it can target foreign organisms and cells while avoiding untoward tissue damage. This is especially important during early pregnancy when the process of implantation might be disrupted. Complement activation is regulated by DAF (also known as CD55), which inactivates the C3 convertase enzymes that activate C3; and by membrane cofactor protein (MCP, also known as CD46), which serves as a cofactor for factor I-­mediated degradation of activated C3 and C4. A third protein, present only in rodents and named compleand MCP-­like activities.

10

246

PART I  The Fundamentals of Reproduction

Crry controls the deposition of activated C3 and C4 on the surface of autologous cells. Fetal demise due to complement deposition and placental inflammation was observed in mice lacking Crry, reflecting unchecked activation of the complement system.542 Complement activation appears to be an essential event in pregnancy loss associated with antiphospholipid syndrome and can be prevented by administration of heparin.543–545 These observations suggest that control of the complement system is essential for fetomaternal tolerance against attack by the innate immune system. In the endometrium, complement component C3, factor B, and DAF are present in the glandular epithelium, and their expression is upregulated in the secretory phase. MCP is expressed in the glandular epithelium throughout the menstrual cycle. Complement receptor type 1 is present in the stroma during the secretory phase, complement receptor 2 is not detectable, and complement receptor 3 is associated with infiltrating leukocytes in the luteal phase. The cyclic changes in C3, factor B, and DAF expression suggest progesterone regulation. In model human endometrial epithelial cell culture systems, however, steroid hormones did not alter DAF expression, but heparin-­binding epidermal-­like growth factor (HB-­EGF) did increase it.166 Thus the steroid hormone effects on expression of some of the complement system proteins may be indirect, perhaps acting through the stromal compartment. Members of the integrin family are now thought to be involved in the regulation of C3, as part of the protective proteins that prevent the cascade of complement activation. DAF or factor H may interact with these cell surface receptors (including the ανβ3 integrin and perhaps CD44) through binding to OPN to limit C3 activation by digesting bound C3b. The ανβ3 integrin, OPN, and DAF all appear synchronously at the time of implantation in human endometrium and may serve as a primary safeguard against complement activation at the time of embryonic attachment, thus avoiding destruction of the early pregnancy by host defenses.

Antimicrobial Peptides Epithelial cells of the female reproductive tract elaborate antimicrobial peptides that presumably guard against ascending infection.546,547 These peptides coat the epithelial surface and enter the cervical mucus. Among the antimicrobial peptides expressed other than complement are lactoferrin, lysozyme, secretory leukocyte protease inhibitor (SLP1), and defensins. The α-­defensins are elaborated by leukocytes and epithelial cells, and the beta-­defensins are by epithelial cells. Defensin-­5 and alpha-­defensin are expressed in the upper half of the stratified squamous epithelium of the vagina and ectocervix, and in columnar epithelial cells of the fallopian tube and endometrium. Beta-­defensin-­1 and -­2 and SLP1 are produced by endometrial glandular epithelial cells. Beta-­defensin-­2 expression is increased by proinflammatory cytokines, and defensin-­5 is also evidently upregulated by inflammation. Defensin-­5 and SLP1 levels in the endometrium also fluctuate during the endometrial cycle, being highest in the secretory phase.

ENDOMETRIAL MICROBIOME The term microbiome refers to a whole set of microorganisms residing within a tissue or organ and includes bacteria, fungi, and viruses. The composition of local microbes can drastically impact human health and disease. Indeed, studies have demonstrated impacts ranging from metabolic to immunologic to epigenetic.548,549 It has been proposed that the endometrial and/or vaginal microbiome plays a role in endometrial receptivity, but the diagnostic criteria and treatment are unclear. At this time, endometrial and vaginal microbiome testing for recurrent implantation failure remains experimental. The microbiome of the female reproductive tract differs from vagina to fallopian tube, and most

bacteria found throughout are lactobacilli (Lactobacillus crispatus, Lactobacillus jensenii, Lactobacillus iners, and Lactobacillus gasseri).550 The presence of Lactobacillus in the vaginal compartment is well established; however, the composition and origin of the uterine cavity or endometrial microbiome are still controversial. In 2016, Moreno and collogues found that high numbers of Lactobacillus are associated with successful implantation, whereas a lower level of Lactobacillus and a high-­level Gardnerella vaginalis and Streptococci can result in decreased reproductive success.551 In contrast, two other studies found no evidence relating Lactobacillus levels and pregnancy success.552,553 It is clear that future research needs to focus on standardizing endometrial sampling for microbiome analysis to experimentally determine its role in reproduction.

CLINICAL EVALUATION OF THE ENDOMETRIUM Endometrial Biopsy The endometrial biopsy has been used to assess endometrial histology for the last 60 to 70 years.554 While sampling the endometrium is used extensively in the setting of abnormal bleeding as a diagnostic tool for endometrial cancer or hyperplasia, its primary role in the evaluation of the infertile couple was, in the past, assessment of an “adequate” luteal phase. However, the clinical utility of the endometrial biopsy as a routine test has been largely abandoned due to the imprecision and inconsistency of histologic dating, which in turn is reflected in the wide diversity of opinion in the literature regarding the incidence and clinical importance of luteal phase deficiency (LPD).555–558 It should be noted, however, that progesterone is essential for the endometrium to become receptive to embryo implantation. The length of progesterone exposure determines when the endometrium becomes receptive.559,560 Thus, below some lower limit of serum progesterone concentration, there must be inadequate receptivity to the embryo. A recent study showed that lowering progesterone levels below about 1.5 ng/mL results in temporal delay in histological markers of endometrial differentiation and levels below about 10 ng/mL resulted in subtle changes in gene expression,561 thus supporting the conceptual underpinnings of luteal phase deficiency. However, these same findings suggest that ovulatory women, whose serum progesterone always exceeds 3 ng/mL, are unlikely to have histological delay solely from low progesterone concentrations. Currently, the best method for diagnosing luteal phase problems may be the presence of a shortened luteal phase, as shown by methods such as basal body temperature charting (Figs. 10.21 and 10.22). While the normal luteal phase should be approximately 12 to 14 days in length (see Fig. 10.23A), anovulatory cycles have no temperature shift (Fig. 10.23B). In the case of LPD, premature luteolysis could potentially compromise the establishment of pregnancy. Such charting can also demonstrate anovulatory cycles (see Fig. 10.23C). However, the endometrial biopsy itself has regained some favor in light of a series of studies that suggest that inflammation associated with disrupting the endometrium and subsequent wound healing results in improved endometrial receptivity for 1 to 3 months afterward.562–565 The benefits of endometrial disruption, the best methods to achieve benefits, and the mechanisms at work remain unclear.566 One mechanistic theory proposed is the accumulation of macrophages and DC after local injury.567–569 Other recent observations regarding luteal function in the endometrium are worthy of consideration. With the completion of the human genome project and the availability of high throughput DNA microarray analyses, the phenomenon of progesterone resistance has been described, based on patterns of gene expression in women with endometriosis compared with fertile controls.121 This phenomenon has been described in ovulatory Whether such changes account

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CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract

for infertility or pregnancy loss associated with these conditions remains to be proven.

that correlate with the putative window of implantation on cycle days 20 to 24 and importance to implantation.575 While certain biomarkers, such as calcitonin or LIF, are aligned closely within this window, others have patterns of expression that are inversely related to the time of maximal endometrial receptivity (epithelial ESR1, PGR, and telomerase). Cell adhesion molecules have been established as well-­ characterized biomarkers of uterine receptivity. L-­ selectin is another cell adhesion molecule noted to be present in the embryo, which binds to the L-­selectin ligand, present on the receptive

Endometrial Receptivity Biomarkers and Clinical Evaluation Attempts continue to find suitable biomarkers of endometrial receptivity.327,570–574 A partial list of putative receptivity biomarkers of receptivity is provided in Table 10.1. Endometrial factors achieve relevance based on their temporal patterns of expression Last 12 cycles Shortest Cycle day Date Day of week Intercourse Birth control Time Temp count

Waking temperature

A

Cycle day Menses

B

Cycle day Menses

Longest

33

Luteal phase

9

Time cycle, Length

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 7 8 9 10 11 12 23 24 25 26 27 28 29 30 31 1 2 3 4 5 6 Fri Sat Sun Mon Tue Wed Thu Fri Sat Sun Mon Tue Wed Thu Fri Sat Sun Mon Tue Wed Thu

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 3 2 1 97 9 8 1 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 2 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 3 4

Cycle day 1 2 3 Date 15 16 17 Day of week Wed Thu Fri Intercourse Birth control Time Temp count

Waking temperature

20

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 1 4

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 2 4

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 3 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 5

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 6

4

4

4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 8

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 9

4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 10

4 5 6 7 8 9 18 19 20 21 22 23 Sat Sun Mon Tue Wed Thu 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 11

10 24 Fri

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 12

9 9 8 8 7 7 6 6 5 5 4 4 3 3 2 2 1 1 98 98 9 9 8 8 7 7 6 6 5 5 4 4 3 3 2 2 1 1 97 97 9 9 8 8 13 14

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 5

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 6

4

4

4

4

9 9 8 8 7 7 6 6 5 5 4 4 3 3 2 2 1 1 98 98 9 9 8 8 7 7 6 6 5 5 4 4 3 3 2 2 1 1 97 97 9 9 8 8 15 16

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 17

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 18

4:00 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 7

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 8

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 9

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 10

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 11

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 12

22 23 24 25 26 27 28 29 30 31 32 13 14 15 16 17 18 19 20 21 22 23 Fri Sat Sun Mon Tue Wed Thu Fri Sat Sun Mon

1 9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 19

17 31 Fri

2 9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 20

3 9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 21

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 22

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 23

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 14

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 15

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 16

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 24

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 25

3:00 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 17

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 26

18 19 20 21 22 23 1 2 3 4 5 6 Sat Sun Mon Tue Wed Thu 4 4 4

4:45 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 13

14

4

16 11 12 13 14 15 30 25 26 27 28 29 Sat Sun Mon Tue Wed Thu

3:45 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 4 4

4

32 Luteal phase

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 18

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 19

3 2 1 98 8 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 20

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 21

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 27

24 7 Fri

4:30 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 22

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 23

Fig. 10.21 Basal body temperature charts can be helpful for the diagnosis of ovulatory dysfunction. Compared with normal 14-day luteal phase (A), anovulatory cycles fail to show a normal thermogenic shift (

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 28

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 29

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 30

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 31

25 26 27 28 8 9 10 11 Sat Sun Mon Tue 4 4 3:30 4:50

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 24

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 32

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 25

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 26

4:01 1 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 27

3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7 6 5 4 3 2 28

10

248

PART I  The Fundamentals of Reproduction Last 12 cycles Shortest Cycle day Date Day of week Intercourse Birth control Time Temp count

Waking temperature

C

Cycle day Menses

1 23 Fri

20

Longest

33

2 3 4 5 6 7 24 25 26 27 28 29 Sat Sun Mon Tue Wed Thu 4

Luteal phase 8 30 Fri 4

9 31 Sat 4

9

Time cycle, Length

10 11 12 13 14 1 2 3 4 5 Sun Mon Tue Wed Thu 4 4 7:30

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 1 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 2 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 3 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 5

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 6

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 7

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 8

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 9

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 10

15 6 Fri 4

16 7 Sat 4

17 18 19 20 21 8 9 10 11 12 Sun Mon Tue Wed Thu 4 4

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 15

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 16

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 17

32 Luteal phase

14

22 13 Fri

23 24 25 26 27 14 15 16 17 18 Sat Sun Mon Tue Wed

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 22

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 23

7:30 9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 11

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 12

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 13

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 14

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 18

1 9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 19

2 9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 20

3 9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 21

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 24

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 25

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 26

9 8 7 6 5 4 3 2 1 98 9 8 7 6 5 4 3 2 1 97 9 8 27

Fig. 10.21, cont’d  In some cases, a short luteal phase is evident (C).

endometrium during the window of implantation.321,337,576,577 Reductions in the L-­selectin ligand occur in certain conditions, including endometriosis and implantation failure.337,578,579 Integrins are cell adhesion molecules noted to be involved in embryo implantation in human and animal models.130,132,300 As shown in Fig. 10.22, three epithelial integrins undergo changes in expression during the luteal phase. This pattern has now been consistently observed in multiple studies.328,580–585 The ανβ3 is regulated directly by HOXA10,586 and these two biomarkers have been extensively studied in the setting of endometriosis, tubal disease, and infertility.38,129,318,530,531,587–590 The ανβ3 integrin is present on the uppermost portion of the luminal epithelium corresponding to the localization of OPN and pinopodes.273,343 Other proteins are also being studied that have relevance to the implantation process, including LIF, and appear related to the decreased integrin expression in women with endometriosis and infertility.591

Global Gene Expression Patterns During the Window of Implantation The dramatic changes in expressed genes regulating growth, secretions, cell growth, and then cell death have been studied at the molecular level using DNA microarray. Demonstration of the key patterns of gene expression between the prereceptive (early secretory) phase and receptive (midsecretory) phase was first defined in 2002; 370 genes (3.1%) displayed decreases ranging from 2-­to greater than 100-­fold, while 323 genes (2.7%) displayed increases ranging from 2-­to greater than 45-­fold.592 A survey of the gene expression patterns has recently been completed in normal fertile women throughout the menstrual cycle.593 The patterns of up-­ and downregulated genes appear to define well the four stages of endometrial development and suggest that the functional characteristics of each phase are largely defined by the proteins that are expressed during each phase. Cell cycle proteins are the predominant feature of proliferating endometrium, while the secretory profile of gene expression characterizes the early secretory phase. Preparation for implantation reflects the changes in the

extracellular matrix, cell communication and adhesion, negative effectors of cell proliferation, synthesis of amino acids, cell ion homeostasis, and appearance of genes involved in innate immunity are all upregulated during this time. Finally, with menstruation, there are dramatic changes in gene expression toward cell death, enzymatic digestive processes and endocytosis/phagocytosis, hemostasis, and antimicrobial responses to the shedding of the endometrium during menstruation. The molecular phenotype of the human endometrium in the midsecretory phase has been characterized using gene analysis of RNA extracted from endometrial biopsies obtained from fertile women.592–595 A relatively small percentage of genes (less than 6%) represented on the chips were significantly increased or reduced in expression (at least twofold) in the presumed receptive phase. The microarray analysis confirmed the upregulation of transcripts previously known to be increased during the window of implantation (including OPN, DAF, glycodelin, and IGFBP-­1) and the downregulation of others (including MMP-­7 and cyclin B). However, several previously unappreciated genes changed in expression during the presumed window of implantation, most notably members of the WNT signaling pathway (see Fig. 10.7A and B). Such changes included the increased expression of the WNT pathway inhibitor DKK1 and a striking decline in the soluble form of FrpHE (SFRP4), another WNT pathway inhibitor. Carson et al also reported a large increase in WNT-­l0b expression and a large decrease in soluble FrpHE (an inhibitor of the WNT pathway), an interesting observation, given existing evidence implicating this signaling molecule in glandular development and hyperplasia in the mammary glands. The microarray analysis also revealed significant alterations in genes encoding water and ion transporters, including upregulation of claudin-­4/Clostridium perfringens enterotoxin receptor; prostaglandin synthesis and action were also found to be altered, including upregulation of phospholipase A2 and prostaglandin E2 receptor (PTGER1 or EP1). The functional significance of these genes, as well as the alterations in others, remains to be determined. Moreover, the interpretation of the findings is complicated by the fact that alterations in specific endometrial cellular

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract

249

4

10

3 1 4

4

3 2 1

HSCORE

4

3

3 2 1

4

1

3 2 1

Menses

Ovulation day 14

Implantation days 20–24

compartments could not be ascertained for most of the genes of interest. The data sets do, however, provide a first approximation of the molecular signature of the human endometrium during the putative time of implantation in a small group of fertile women. This information has been used to develop a diagnostic “chip” profile for establishing changes in the timing of the fertile window, although the clinical utility of these approaches remains to be fully tested.596–597 More recently, single-­cell RNA sequencing has provided evidence of the shifting pattern of cell types in the human endometrium through the menstrual cycle598 and allows detection of changes in RNA expression from each cell type, without being lost in combination with others. One finding of interest is that there is an abrupt change in the transcriptome of unciliated epithelial cells at the time the endometrium becomes receptive, while late secretory changes are more continuous, consistent with the rare but observed later implantations. Of note, with these efforts to identify the endometrial transcriptome across the entire menstrual cycle, investigators can now view each segment of the menstrual cycle as a unique fingerprint detailing the role of steroid hormones and their receptors and the endometrium.593 The genes that are expressed during the proliferative phase in response to estrogen play roles in cell adhesion, cell-­cell signaling, cell cycle regulation, and cell division. After ovulation in the early secretory phase in response to both ovarian estrogen and progesterone, gene expression shifts globally to include metabolic enzymes, transporter proteins, inhibitors of WNT signaling with a marked upregulation of lipid metab olism, phospholipase activity, and prostaglandin metabolism.

Pregnancy

Fig. 10.22 Scattergram showing distribution of three cycle-­dependent epithelial integrin subunits in cycling endometrium.  While each of the integrin subunits has a unique pattern of expression, all three are coexpressed during the time of maximal receptivity between cycle days 20 and 24. (From Lessey BA. Two pathways of progesterone action in the human endometrium: implications for implantation and contraception. Steroids. 2003;68:809–815.)

There is increased EGF signaling noted during this time. The midsecretory is distinct from the other stages due to the downregulation of ESR1 and PGR in the epithelial compartment, and this is reflected in the gene expression profiling. With this loss of estrogen action and dominance of progesterone action in the stroma comes a shift in cell adhesion, communication, and motility. There is an increase in intracellular signaling, antiapoptosis, and immune response, including DAF, defensin, complement component 4, and glycodelin. These detailed DNA microarray assays are providing the basis for new biopsy-­based assessment of endometrial progression and are being promoted as assessment tools to improve IVF outcomes.599,600 During the late secretory phase, progesterone withdrawal leads to preparation for menstruation with increased expression of MMPs, EBAF (LEFTY; an anti-­TGFB cytokine), bone morphogenetic protein-­2, and signaling through the IL2 receptor. Evidence of leukocyte infiltration appears with gene expression patterns reflecting leukocyte integrins. Genes encoding proteins for the humoral immune response are further upregulated at this time, while the most downregulated reflect a passing window of receptivity, with loss of LIF, DKK1, IGF1, and complement proteins. Thus, each of the four segments of the menstrual cycle have now been well-­defined using DNA microarray technology, integrating our understanding of how shifts in ovarian steroids and their receptors influence the different phases of the menstrual cycle. The pattern of gene expression during the midsecretory phase has given way to the development of gene patterns as biomarkers of endometrial differentiation, proposed as an improvement

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PART I  The Fundamentals of Reproduction

Endometriosis

P4 Resistance

E2 Dominance

Progesterone COX2

INF , TNF , IL-1, Il-17 SIRT1 BCL6 COUP-TFII, HAND 2, HOXA10, HBEGF, BMP2, FGF, 17HSDII, STRA6, CRABPII, FOX01

PGE2 IL-6 and IL-10

PIAS3 P450arom

pSTAT3

HIF1A E2

Inflammation Angiogenesis Pregnancy

Immunosuppression

VEGF, CYR61, Haptoglobin, Ang-2, c-Met Adrenomedulli

Cell Proliferation Fig. 10.23 Like menstruation, endometriosis is associated with inflammation.  Inflammatory cytokines including interferon-­gamma (INFγ), tumor necrosis factor α (TNFα), interleukin (IL)-­1, and IL-­17 have downstream effects on the endometrium. IL-­17 has been shown to stimulate cyclooxygenase 2 (COX2) expression and prostaglandin (PGE2) production that stimulates aromatase expression. Endometrial IL-­6 is associated with these changes that activate signal transducer and activator of transcription-­3 (STAT3). Phosphorylated STAT3 (pSTAT3), as seen in endometriosis and infertility,104 stabilizes hypoxia-­induced factor-­ 1α (HIF-­1α), which together with STAT3 stimulates many of the estrogen-­dependent changes associated with endometriosis, including estrogen receptor β. STAT3 also induces BCL6, which is a candidate signaling protein that is associated with endometriosis and progesterone resistance.112 The imbalance between estrogen and progesterone actions likely plays a critical role in the implantation defects found associated with various inflammatory states including endometriosis, adenomyosis, and hydrosalpinges. Ang-­2, Angiopoietin; BMP2, bone morphogenic protein 2; c-­MET, hepatocyte growth factor receptor; COUP-­TFII, chicken ovalbumin protein transcription factor II; CRABPII, cellular retinoic acid binding-­protein 2; CYR61, cysteine-­rich angiogenic inducer 61; E2, estradiol; FGF, fibroblast growth factor; FOXO1, forkhead box protein O1; HAND2, heart and neural crest derivatives expressed protein-­2; HBEGF, heparin-­binding epidermal growth factor-­like growth factor; HOXA10, homeobox A10; P450arom, P450 aromatase; PIAS3, protein inhibitor of activated STAT3; STRA6, stimulated by retinoic acid 6; VEGF, vascular endothelial growth factor.

over histology.601–603 Based on the ability to define the window of implantation, these assays are now designed to help better establish and precisely define the time of embryo transfer in IVF.597,600 One such test, the Endometrial Receptivity Array (ERA) consists of a customized microarray based on the transcriptomic signature of human receptive endometrium, specifically when the endometrium is receptive to blastocyst adhesion and attachment.299,604,664 Thus, the ERA test is employed to determine the optimal timing of the window of implantation for embryo transfer in a clinical setting. The ERA test divides the results of the endometrial biopsy into receptive and nonreceptive, and further to prereceptive or postreceptive to better synchronize the embryo and the uterus.605,606,607 Despite the promising early results from a small cohort of women, the ERA test has not shown any clinical advantage for patients that had failed three or more embryo transfers.608

Inflammatory Conditions and the Endometrium and Reproductive Tract Endometriosis is an inflammatory disease, arising from immune activation in response to ectopic endometrium (see Chapter 25),609 creating an autoimmune phenomenon that targets the

eutopic endometrium.610 As recently reviewed by Taylor et al,611 alterations in chemokines may directly target T cells, monocytes, and other leukocytes, including MCP-­1 (CCL2), MIP-­1 alpha (CCL3), Rantes (CCL5), and eotaxin (CCL11). Other chemokines attract monocytes and neutrophils, including Gro-­a (CXCL1), epithelial cell-­ derived neutrophil-­ activating peptide (ENA-­ 78; CXCL-­ 5), and stromal cell-­ derived factor (SDF-­ 1; CXCL12). A new theory has been promoted that progesterone resistance and therefore estrogen dominance arises from the resulting inflammatory response,612 activating factors that perturb normal progesterone signaling through the IHH pathway (see Fig. 10.4).613 In normal endometrium, progestins act through PGR, which is dependent on epithelial transcription factor GATA-­2.295 PGR induces epithelial IHH that signals via paracrine mechanisms to the underlying stroma to initiate COUP-­TFII with the help of signaling molecule, GLI1, to suppress estrogen action inflammation.614–616 In inflammatory conditions such as endometriosis or hydrosalpinx, interleukin 17 (IL17) and IL6 are systemically or locally increased.617–620 As shown in Fig. 10.23, elevated IL6 triggers phosphorylation and activation of STAT3, with stabiThis molecule is normally not seen until

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract

251

TABLE 10.1  Endometrial Biomarkers Used for Assessment of Endometrial Receptivity

10

Functional Category

Biomarker

Expression Pattern

Comments

Chemokine/cytokine

LIF IL1,11 17HSD-II Aromatase

m----------o--- ------- -m m----------o-- ------- --m m----------o-- --------- m m----------o-----------m

All essential for implantation

Enzymes Extracellular matrix/ receptors/ligands

Growth factors

Major secreted proteins Paracrine or autocrine Steroid receptors and related cofactors

Transcription factors

Miscellaneous

Integrins 1 1 4 4 3 MMPs EBAF (lefty) MUC-1 L-selectin ligand Cadherin-11 TGF EGF Ligand family Hepatocyte growth factor (HGF) Heparin binding-EGF-like growth factor (HB-EGF) IGF-I IGF-II Glycodelin Calcitonin Osteopontin Indian hedgehog (IHH) Bone morphogenic protein 2 (BMP2/TGFb1) ER PR-A/PR-B FKBP51, FKPB52 MAGE-11 HOXA10 HOXA11 CCAAT enhancer binding protein(C/EBP ) Forkhead box O1 (FOXO1) Beta catenin CD44 Prolactin

menstruation, a potent inducer of angiogenesis. Mechanisms to deactivate STAT3 are repressed under conditions of inflammation,622 and prolonged STAT3 expression stimulates overexpression of the oncogene B-­cell lymphoma protein 6 (BCL6).623 It has been proposed that BCL6 combines with the histone deacetylase SIRT1 to interfere with expression of GLI1,624 a critical component in the IHH pathway. Exaggerated estrogen responses are also a response to inflammation, including a shift toward PTGS2 and prostaglandin production, which further increases estrogen through aberrant aromatase expression, as seen in endometriosis.625 Thus recent advances in the field of progesterone resistance provide a new understanding of the global changes associated with endometriosis and promise to keep attention focused on endometrial receptivity as a primary cause of infertility.

Adenomyosis Adenomyosis is a common gynecologic disorder seen in women of reproductive age and is defined by the presence of endometrial glands and stroma within the myometrial compartment of the uterus.626,627 It is found in 10% to 66% of women at the time of hysterectomy628 and is a major cause of infertility, pelvic pain, and AUB.629–631 Like endometriosis, adenomyosis is estrogen-­ dependent for its growth.627 WNT-CTNNB1 activation through estrogen-­EGF-­mediated pathways is thought to contribute to the

m---------- o-----------m m---------- o--------- --m m----------o----- ------m m----------o--------- --m m----------o--------- --m m----------o- ---------- m m----------o--- ------- -m m----------o---- ------- m m----------o- --------- -m m----------o----------- m m----------o------ -----m m----------o-- --------- m m---------- o-----------m m----------o- ---------- m m----------o---- ------- m m----------o----- ------m m----------o--- -------- m m----------o- --------- -m m----------o--- -------- m

Converts E2 to less active E1; Should not be present in normal endometrium These 3 integrins frame the window of implantation; MMPs and lefty (EBAF) come up at menses; Epithelial glycoprotein Binds L-selection; Stromal marker Mediates P action; Involved in both proliferation and implantation; Involved in decidualization Stromal and epithelial expression; Proliferative phase Secretory phase All 3 are epithelial proteins Induced by P; Induced by IHH

m----------o-- ---------m m----------o----------- m m----------o- --------- -m m----------o-- -------- -m

Estrogen induced and P inhibited; PRA persists in decidua Each induced by P; Expressed in midsecretory phase

m----------o- --------- -m m----------o- --------- -m m----------o- --------- -m

Both essential for implantation; Expressed during WOI Essential for decidualization; Expressed during proliferation

m----------o- --------- -m m----------o- ----------m m----------o----- ------m m----------o--- -------- m

During WOI; Marker of decidualization

pathogenesis of adenomyosis through epithelial-­ mesenchymal transformation.632 Risk factors for developing adenomyosis include increased parity, as well as prior cesarean delivery.633,634 Adenomyosis is a challenging condition to treat. Medical options include oral contraceptives, long-­ term progestin therapies, gonadotropin-­ releasing hormone (GnRH) agonist therapy, and selective PGR modulators.635 Surgical options remain the definitive management strategy.

Inflammation and MicroRNAs MicroRNAs are small nonprotein coding RNAs that regulate specific posttranscriptional messenger RNAs, targeting them for degradation. Genes coding for miRNA represent 1% to 4% of the genes in higher organisms and are highly processed sequences that are derived from larger RNA transcripts. These small precursor miRNAs of 70 to 90 nucleotides are transported to the cytoplasm, where they undergo cleavage by the RNAse III enzyme, Dicer, to generate a double-­stranded miRNA duplex, eventually forming a single-­stranded miRNA that binds to complementary sequences on target mRNA and preventing their transcription.636–638 This extraordinarily complex regulatory mechanism of transcription has now been studied in the human endometrium characterizaIn endometriosis,

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PART I  The Fundamentals of Reproduction

complex networks of genes appear to be altered on the basis of specific changes in miRNAs,123,124,641–644 providing a new dimension to the study of endometrial receptivity.572 Examination of miRNAs in serum or blood may also provide opportunities for diagnosis of endometriosis.645,646 Specific changes in miRNAs have been noted that appear to directly relate to the pathophysiology of endometriosis and the associated inflammation and progesterone resistance associated with this disease. Elevated miR-­210, for example, has been implicated in STAT3 activation,647 which is involved in proliferation and VEGF production, HIF1A activation, and stimulation of BCL6, implicated in progesterone resistance. Other miRNAs implicated in progesterone resistance include miRNA196a, which is overexpressed in endometriosis and thought to directly downregulate PGR.648 MiRNA-­29c targets the PGR chaperone protein FKPB4, involved in progesterone action. Tissue levels of the miRNA are increased in women and animal models of endometriosis.125 MiRNA-­15a-­5p has been found to be reduced in endometriosis, its target being VEGFA; reduced expression of the miRNA may promote angiogenesis in endometriosis.649

Ultrasonography Abnormalities of the structure of the uterus from congenital defects, neoplasia, or synechiae can impair fertility (see Chapter 25). Diagnostic imaging methods play an important role in the assessment of these uterine defects.650 Transvaginal sonography (TVS) has become widely accepted as a tool for high-­resolution imaging of the female internal reproductive organs.651 This noninvasive, convenient, and safe technology provides rapid diagnoses with a high correlation with pathology. TVS is used primarily to monitor follicular development and endometrial thickness during exogenous hormone treatments of infertile patients. The method might offer a significant advantage for use as a diagnostic aid in various other endometrial conditions, such as endometrial polyps, submucous and intramural leiomyomata, endometrial hyperplasia, and carcinoma. TVS has certainly found its use in the assessment of early pregnancy. Growth of the endometrium can easily be measured using ultrasound. Endometrial thickness and texture are commonly assessed, especially when women are sequentially monitored as part of their treatment with human menopausal gonadotropins. Endometrium in the early proliferative phase immediately following menses is typically thin; in response to estrogen, the endometrium thickens and becomes trilaminar in appearance, growing between 0.1 and 0.5 mm daily. Following ovulation, the endometrium becomes hyperechoic as secretory changes ensue. Various attempts have been made to classify these patterns on the basis of thickness and texture. Most authors suggest that a thickness of 8 mm or greater with a trilaminar appearance is adequate for implantation in an IVF cycle.652–656 Beyond a certain threshold, however, there is no correlation between implantation rates and endometrial thickness. The cyclic endometrial changes induced by varying estrogen and progesterone levels result in predictable sonographic changes, especially in blood flow and vessel density. Both endometrial thickness and echogenic pattern have been studied as potential markers of uterine receptivity and predictors of successful embryo implantation. Transvaginal pulsed Doppler ultrasound measures uterine artery blood flow (or the impedance to flow) and is expressed as the pulsatility index (PI). The PI varies across the menstrual cycle and may be an additional index to predict implantation after assisted reproductive techniques. Aside from endometrial thickness, morphology, blood flow, and uterine artery pulsatility have all been examined as possible markers of a receptive endometrium. Increased PI has been associated with elevated markers of pregnancy loss, including anticardiolipin

antibodies.657,658 Studies in pregnancy using Doppler flow ultrasound and pregnancy rates have been inconclusive.659

Sonohysterography Identification of uterine fibroids, endometriomas, and uterine septa is now routine (see Chapter 25). Lesions such as polyps within the uterine cavity may be misinterpreted as thickened endometrium and therefore go undiagnosed. Instillation of sterile saline into the uterine cavity as part of sonohysterography provides an enhanced view of the uterine cavity and may detect even small lesions. This approach is valuable in assessing the effects of selective estrogen receptor modulators like tamoxifen on the endometrium.660 A sonohysterogram in a normal uterus shows a smooth contour (Fig. 10.24A). In some cases, abnormalities can be more clearly seen with sterile saline installation (see Fig. 10.24A). Three-­ dimensional (3D) sonohysterography can provide a dramatic 3D rendering of the uterine cavity, giving a better overall appreciation of such lesions, and better determining the location and point of attachment (see Fig. 10.24C).661 This evolving technology may further improve the sensitivity and specificity of the technique used to detect small polyps or fibroids. With the advent of 3D ultrasound, differential diagnoses between a septate versus a bicornuate uterus can also be more readily determined, avoiding the need for more expensive modalities like MRI. The use of 3D ultrasound for the assessment of uterine receptivity is also evolving.662

Hysteroscopy Hysteroscopy is a highly useful tool to identify and correct lesions identified by hysterosalpingography or sonohysterography (see Fig. 10.24D).663,664 Visualization of the endometrial cavity has proven useful for the inspection of the uterine cavity in women with abnormal bleeding, infertility, and RPL. Direct visualization allows the operator to resect or biopsy lesions once identified. Resection of uterine septa in müllerian fusion defects is commonly performed. More recently, sterilization techniques through the hysteroscopic have been refined to a point that challenges other techniques and are readily available in many centers around the world.665

Endometrial Bleeding The endometrium is the only tissue in the body that has evolved the capacity to bleed on a regular basis and stop bleeding and undergo repair after each menstrual cycle. Bleeding episodes outside of menses are a frequent occurrence at the boundaries of reproductive life, occurring more frequently at menarche and at perimenopause. Evaluation begins with an assessment of the reproductive status of the individual. A thorough history and physical, as well as laboratory and imaging studies, are required to make a proper diagnosis. In the absence of ovulation, and without progesterone, the endometrium proliferates under the influence of estrogen and can break down, resulting in prolonged and irregular bleeding episodes. In the absence of uterine pathologies such as endometrial polyps, fibroids, endometrial hyperplasia or neoplasia, or coagulopathy, anovulatory bleeding is the most frequent diagnosis (see Chapter 25). hCG testing can be performed to confirm pregnancy, and ultrasound and endometrial biopsy can be performed to rule out endometritis, hyperplasia, or endometrial malignancy. Saline infusion sonography or hysteroscopy is helpful in the diagnosis of anatomic abnormalities, including fibroids, retained products of conception, or endometrial polyps. Bleeding associated with trauma or sexually transmitted infections should also be considered. Von Willebrand disease is the most frequent coagulopathy, especially in young women.666 AUB is an increasingly common complaint, frequently associated with ovulatory dysfunction and presenting with heavy and

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10

A

B

C

Fig. 10.24 Sonohysterography can be used to visualize intracavitary lesions.  Injection of sterile saline shows the uterine cavity to be free of polyps or fibroids (A). In the case of intrauterine pathology, lesions such as a polyp can be easily delineated either in 2D (B) or 3D rendering (C). Hysteroscopy confirms the polyp (D), which can then be easily resected.

D

irregular occurrence of bleeding.667 Anovulatory bleeding is often painless, without the usual uterine cramping experienced during normal menstruation. AUB is frequently associated with obesity and insulin resistance and results from anovulation and is associated with PCOS.668,669 Findings of polycystic ovaries are common and part of the diagnostic triad of making the diagnosis of PCOS.670 In chronic PCOS with prolonged amenorrhea with irregular bleeding, endometrial pathology can develop, resulting in hyperplasia or malignancy.671 While this occurrence is rare in young women (0.2 per 100,000 cases), the incidence increases to 14 to 24 per 100,000 in those after age 40.667 Treatment of AUB is directed at the cause. While surgery may be required for anatomic pathology, medical therapy is the standard of care for most AUB. In the case of anovulatory bleeding, medical treatment with oral contraceptives or continuous or intermittent progestins is the first line of treatment, despite no randomized trials supporting this approach. Long-­acting reversible contraception (LARC) has proven helpful for the treatment of anovulatory bleeding, as has weight loss and lifestyle changes.672

ENDOMETRIUM IN ADVANCING AGE Based on donor oocyte pregnancies achieved in postmenopausal women whose reproductive tract was programmed with exogenous steroid hormones, uterine aging in the absence of acquired pathology evidently does not preclude successful implantation and carriage of a pregnancy to term. The relentless ability of the endometrium to restore itself may hint at its longevity. A rich source of the enzyme telomerase,197 the endometrium may have the ability to delay the normal aging process. However, there are structural changes that take place in the uterus with advancing age, pregnancy, and the cessation of ovarian function. Whether or not these changes affect the “reproductive potential” of the uterus or contribute to the increased incidence of pregnancy complications associated with advanced maternal age is not known. The basal endometrium interdigitates with the myometrium with advancing age, resulting in a degree of superficial adeno myosis that is a normal finding in the uterus in the fifth decade

of life. The infiltrating endometrium does not undergo normal cyclic changes. This may be a consequence of a uterus hosting a past pregnancy.673 After menopause, in the absence of hormone replacement, endometrial atrophy is apparent and mitotic activity ceases. Epithelial cells shrink in size, and the stroma becomes fibrotic. A compact eosinophilic material is found in the lumina of the endometrial glands, occasionally engorging them and giving rise to the histologic pattern referred to as cystic atrophy. TOP REFERENCES

Cha J, Sun X, Dey SK. Mechanisms of implantation: strategies for successful pregnancy. Nat Med. 2012;18:1754–1767. Fox C, Morin S, Jeong JW, et al. Local and systemic factors and implantation: what is the evidence? Fertil Steril. 2016;105:873–884. Gellersen B, Brosens IA, Brosens JJ. Decidualization of the human endometrium: mechanisms, functions, and clinical perspectives. Semin Reprod Med. 2007;25(6):445–453. Johannesson L, Jarvholm S. Uterus transplantation: current progress and future prospects. Int J Womens Health. 2016;8:43–51. Kelleher AM, Demayo FJ, Spencer TE. Uterine Glands: Developmental Biology and Functional Roles in Pregnancy. Endocr Rev. 2019;40(5):1424–1445. Large MJ, Demayo FJ. The regulation of embryo implantation and endometrial decidualization by progesterone receptor signaling. Mol Cell Endocrinol. 2012;358:155–165. May KE, Villar J, Kirtley S, et al. Endometrial alterations in endometriosis: a systematic review of putative biomarkers. Hum Reprod Update. 2011;17:637–653. Palomba S, Homburg R, Santagni S, et al. Risk of adverse pregnancy and perinatal outcomes after high technology infertility treatment: a comprehensive systematic review. Reprod Biol Endocrinol. 2016;14:76. Robboy SJ, Kurita T, Baskin L, Cunha GR. New insights into human female reproductive tract development. Differentiation. 2017;97:9–22. Salamonsen LA, Evans J, Nguyen HP, et al. The microenvironment of human implantation: determinant of reproductive success. Am J Reprod Immunol. 2016;75:218–225.

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REFERENCES

1. Kobayashi A, Behringer RR. Developmental genetics of the female reproductive tract in mammals. Nat Rev Genet. 2003;4(12):969–980. 2. Orvis GD, Behringer RR. Cellular mechanisms of Müllerian duct formation in the mouse. Dev Biol. 2007;306(2):493–504. 3. O’Rahilly R. Prenatal human development. In: Wynn RM, Jolie WP, eds. Biology of the Uterus. Plenum Publishing; 1989:35. 4. Szamborski J, Laskowska H. Some observations on the developmental histology of the human foetal uterus. Biol Neonat. 1968;13(5):298–314. 5. Gray CA, Bartol FF, Tarleton BJ, et al. Developmental biology of uterine glands. Biol Reprod. 2001;65(5):1311–1323. 6. Kurita T. Normal and abnormal epithelial differentiation in the female reproductive tract. Differentiation. 2011;82(3):117–126. 7. Spencer TE, Dunlap KA, Filant J. Comparative developmental biology of the uterus: insights into mechanisms and developmental disruption. Mol Cell Endocrinol. 2012;354(1–2):34–53. 8. Cunha GR, Kurita T, Cao M, Shen J, Robboy S, Baskin L. Molecular mechanisms of development of the human fetal female reproductive tract. Differentiation. 2017;97:54–72. 9. Robboy SJ, Kurita T, Baskin L, Cunha GR. New insights into human female reproductive tract development. Differentiation. 2017;97:9–22. 10. Heikkila M, Peltoketo H, Vainio S. Wants and the female reproductive system. J Exp Zool. 2001:616–623. Published online. 11. Taylor HS, Heuvel V, Igarashi GB, Igarashi P. A conserved Hox Axis in the mouse and human female reproductive system: late establishment and persistent adult expression of the Hoxa cluster genes. Biol Reprod. 1997;57:1338–1345. 12. Wu HH, Reindollar RH, Gray MR. Analysis of the WNT-­4 gene in women with congenital absence of the uterus and vagina. Fertil Steril. 2002;78:S169–S170. 13. Bartol FF, Wiley AA, Bagnell CA. Uterine development and endometrial programming. Soc Reprod Fertil Suppl. 2006;62:113–130. 14. Kobayashi A, Shawlot W, Kania A, Behringer RR. Requirement of Lim1 for female reproductive tract development. Development. 2004;131(3):539–549. 15. Torban E, Dziarmaga A, Iglesias D, et al. PAX2 activates WNT4 expression during mammalian kidney development. J Biol Chem. 2006;281(18):12705–12712. 16. Daftary GS, Taylor HS. EMX2 gene expression in the female reproductive tract and aberrant expression in the endometrium of patients with endometriosis. J Clin Endocrinol Metab. 2004;89(5):2390–2396. 17. Taylor HS, Fei X. Emx2 regulates mammalian reproduction by altering endometrial cell proliferation. Mol Endocrinol. 2005;19(11):2839–2846. 18. Miller C, Pavlova A, Sassoon DA. Differential expression patterns of Wnt genes in the murine female reproductive tract during development and the estrous cycle. Mech Dev. 1998;76(1–2):91–99. 19. Franco HL, Dai D, Lee KY, et al. WNT4 is a key regulator of normal postnatal uterine development and progesterone signaling during embryo implantation and decidualization in the mouse. FASEB J. 2011;25(4):1176–1187. 20. Carroll TJ, Park JS, Hayashi S, Majumdar A, McMahon AP. Wnt9b plays a central role in the regulation of mesenchymal to epithelial transitions underlying organogenesis of the mammalian urogenital system. Dev Cell. 2005;9(2):283–292. 21. Waschk DEJ, Tewes AC, Römer T, et al. Mutations in WNT9B are associated with Mayer-­ Rokitansky-­ Küster-­ Hauser syndrome. Clin Genet. 2016;89(5):590–596. 22. Ma W, Li Y, Wang M, et al. Associations of polymorphisms in WNT9B and PBX1 with Mayer-­Rokitansky-­Küster-­Hauser syndrome in Chinese Han. PLoS One. 2015;10(6):e0130202. 23. Pizzo A, Laganà AS, Sturlese E, et al. Mayer-­Rokitansky-­Kuster-­ Hauser syndrome: embryology, genetics and clinical and surgical treatment. ISRN Obstet Gynecol. 2013;2013:628717. 24. Morcel K, Dallapiccola B, Pasquier L, Watrin T, Bernardini L, Guerrier D. Clinical utility gene card for: Mayer–Rokitansky– Küster–Hauser syndrome. Eur J Hum Genet. 2012;20(2):1–3. 25. Sandbacka M, Laivuori H, Freitas É, et al. TBX6, LHX1 and copy number variations in the complex genetics of Müllerian aplasia. Orphanet J Rare Dis. 2013;8(1):125. 26. Williams LS, Kim HG, Kalscheuer VM, et al. A balanced chromosomal translocation involving chromosomes 3 and 16 in a patient

















with Mayer-­Rokitansky-­Kuster-­Hauser syndrome reveals new candidate genes at 3p22.3 and 16p13.3. Mol Cytogenet. 2016;9(1):57. 27. Miller C, Sassoon DA. Wnt-­7a maintains appropriate uterine patterning during the development of the mouse female reproductive tract. Development. 1998;125(16):3201–3211. 28. Mericskay M, Kitajewski J, Sassoon D. Wnt5a is required for proper epithelial-­ mesenchymal interactions in the uterus. Development. 2004;131(9):2061–2072. 29. Ravel C, Lorenço D, Dessolle L, et al. Mutational analysis of the WNT gene family in women with Mayer-­ Rokitansky-­ Kuster-­ Hauser syndrome. Fertil Steril. 2009;91(suppl 4):1604–1607. 30. Timmreck LS, Pan HA, Reindollar RH, Gray MR. WNT7A Mutations in patients with Müllerian duct abnormalities. J Pediatr Adolesc Gynecol. 2003;16(4):217–221. 31. Deutscher E, Hung-­Chang Yao H. Essential roles of mesenchyme-­ derived beta-­catenin in mouse Müllerian duct morphogenesis. Dev Biol. 2007;307(2):227–236. 32. Couse JF, Hewitt SC, Bunch DO, et al. Postnatal sex reversal of the ovaries in mice lacking estrogen receptors α and β. Science. 1999;286(5448):2328–2331. 33. Greco TL, Duello TM, Gorski J. Estrogen receptors, estradiol, and diethylstilbestrol in early development: the mouse as a model for the study of estrogen receptors and estrogen sensitivity in embryonic development of male and female reproductive tracts. Endocr Rev. 1993;14(1):59–71. 34. Satokata I, Benson G, Maas R. Sexually dimorphic sterility phenotypes in Hoxa10-­deficient mice. Nature. 1995;374(6521):460–463. 35. Goodman FR, Bacchelli C, Brady AF, et al. Novel HOXA13 mutations and the phenotypic spectrum of hand-­foot-­genital syndrome. Am J Hum Genet. 2000;67(1):197–202. 36. Innis JW, Goodman FR, Bacchelli C, et al. A HOXA13 allele with a missense mutation in the homeobox and a dinucleotide deletion in the promoter underlies Guttmacher syndrome. Hum Mutat. 2002;19(5):573–574. 37. Taylor HS, Igarashi P, Olive DL, Arici A. Sex steroids mediate HOXA11 expression in the human peri-­implantation endometrium. J Clin Endocrinol Metab. 1999;84(3):1129–1135. 38. Taylor HS, Arici A, Olive D, Igarashi P. HOXA10 is expressed in response to sex steroids at the time of implantation in the human endometrium. J Clin Invest. 1998;101(7):1379–1384. 39. Saravelos SH, Cocksedge KA, Li TC. Prevalence and diagnosis of congenital uterine anomalies in women with reproductive failure: a critical appraisal. Hum Reprod Update. 2008;14(5):415–429. 40. Li S, Qayyum A, Coakley FV, Hricak H. Association of renal agenesis and mullerian duct anomalies. J Comput Assist Tomogr. 2000;24(6):829–834. 41. Press MF, Udove JA, Greene GL. Progesterone receptor distribution in the human endometrium. Analysis using monoclonal antibodies to the human progesterone receptor. Am J Pathol. 1988;131(1):112–124. 42. Press MF, Nousek-­ Goebl NA, Bur M, Greene GL. Estrogen receptor localization in the female genital tract. Am J Pathol. 1986;123(2):280–292. 43. Lessey BA, Killam AP, Metzger DA, Haney AF, Greene GL, McCarty Jr KS. Immunohistochemical analysis of human uterine estrogen and progesterone receptors throughout the menstrual cycle. J Clin Endocrinol Metab. 1988;67(2):334–340. 44. Brenner RM, West NB, McClellan MC. Estrogen and progestin receptors in the reproductive tract of male and female primates. Biol Reprod. 1990;42(1):11–19. 45. Gregory CW, Wilson EM, Apparao KBC, et al. Steroid receptor coactivator expression throughout the menstrual cycle in normal and abnormal endometrium. J Clin Endocrinol Metab. 2002;87(6):2960–2966. 46. Han SJ, DeMayo FJ, Xu J, Tsai SY, Tsai MJ, O’Malley BW. Steroid receptor coactivator (SRC)-­1 and SRC-­3 differentially modulate tissue-­ specific activation functions of the progesterone receptor. Mol Endocrinol. 2006;20(1):45–55. 47. Szwarc MM, Kommagani R, Lessey BA, Lydon JP. The p160/steroid receptor coactivator family: potent arbiters of uterine physiology and dysfunction. Biol Reprod. 2014;91(5):122. 48. Clark JH, Gorski J. Ontogeny of the estrogen receptor during early uterine development. Science. 1970;169(3940):76–78. 49. Ogasawara Y, Okamoto S, Kitamura Y, Matsumoto K. Proliferative pattern of uterine cells from birth to adulthood in intact, neonatally castrated, and/or adrenalectomized mice, assayed by incorporation . 1983;113(2):582–587.

253.e1

253.e2 PART I  The Fundamentals of Reproduction 50. Branham WS, Sheehan DM. Ovarian and adrenal contributions to postnatal growth and differentiation of the rat uterus. Biol Reprod. 1995;53(4):863–872. 51. Revankar CM, Cimino DF, Sklar LA, Arterburn JB, Prossnitz ER. A transmembrane intracellular estrogen receptor mediates rapid cell signaling. Science. 2005;307(5715):1625–1630. 52. Spencer TE, Kelleher AM, Bartol FF. Development and function of uterine glands in domestic animals. Annu Rev Anim Biosci. 2019;7(1):125–147. 53. Hewitt SC, Deroo BJ, Korach KS. Signal transduction. A new mediator for an old hormone? Science. 2005;307(5715):1572–1573. 54. Akingbemi BT, Ge R, Rosenfeld CS, et al. Estrogen receptor-­ alpha gene deficiency enhances androgen biosynthesis in the mouse Leydig cell. Endocrinology. 2003;144(1):84–93. 55. Key TJ, Pike MC. The dose-­effect relationship between “unopposed” oestrogens and endometrial mitotic rate: its central role in explaining and predicting endometrial cancer risk. Br J Cancer. 1988;57(2):205–212. 56. Conneely OM, Mulac-­Jericevic B, DeMayo F, Lydon JP, O’Malley BW. Reproductive functions of progesterone receptors. Recent Prog Horm Res. 2002;57(1):339–355. 57. Lee DK, Kurihara I, Jeong JW, et al. Suppression of ERalpha activity by COUP-­ TFII is essential for successful implantation and decidualization. Mol Endocrinol. 2010;24(5):930–940. 58. Tan J, Paria BC, Dey SK, Das SK. Differential uterine expression of estrogen and progesterone receptors correlates with uterine preparation for implantation and decidualization in the mouse. Endocrinology. 1999;140(11):5310–5321. 59. Lessey BA, Palomino WA, Apparao KBC, Young SL, Lininger RA. Estrogen receptor-­alpha (ER-­alpha) and defects in uterine receptivity in women. Reprod Biol Endocrinol. 2006;4(suppl 1):S9. 60. Young SL, Lessey BA. Progesterone function in human endometrium: clinical perspectives. Semin Reprod Med. 2010;28(1):5–16. 61. Hapangama DK, Kamal AM, Bulmer JN. Estrogen receptor beta: the guardian of the endometrium. Hum Reprod Update. 2015;21:174–193. 62. Dupont S, Krust A, Gansmuller A, Dierich A, Chambon P, Mark M. Effect of single and compound knockouts of estrogen receptors alpha (ERalpha) and beta (ERbeta) on mouse reproductive phenotypes. Development. 2000;127(19):4277–4291. 63. Wada-­Hiraike O, Hiraike H, Okinaga H, et al. Role of estrogen receptor beta in uterine stroma and epithelium: insights from estrogen receptor beta-­/-­mice. Proc Natl Acad Sci U S A. 2006;103(48):18350–18355. 64. Frasor J, Barnett DH, Danes JM, Hess R, Parlow AF, Katzenellenbogen BS. Response-­specific and ligand dose-­dependent modulation of estrogen receptor (ER) alpha activity by ERbeta in the uterus. Endocrinology. 2003;144(7):3159–3166. 65. Winuthayanon W, Hewitt SC, Orvis GD. Uterine epithelial estrogen receptor alpha is dispensable for proliferation but essential for complete biological and biochemical responses. Proc Natl Acad Sci U S A. 2010;107:19272–19277. 66. Han SJ, Jung SY, Wu SP. Estrogen receptor beta modulates apoptosis complexes and the inflammasome to drive the pathogenesis of endometriosis. Cell. 2015;163:960–974. 67. Critchley HOD, Henderson TA, Kelly RW, et al. Wild-­type estrogen receptor (ERbeta1) and the splice variant (ERbetacx/beta2) are both expressed within the human endometrium throughout the normal menstrual cycle. J Clin Endocrinol Metab. 2002;87(11):5265–5273. 68. Bombail V, Gibson DA, Collins F, MacPherson S, Critchley HOD, Saunders PTK. A Role for the orphan nuclear receptor estrogen-­ related receptor alpha in endometrial stromal cell decidualization and expression of genes implicated in energy metabolism. J Clin Endocrinol Metab. 2010;95(10):E224–E228. 69. Bombail V, MacPherson S, Critchley HOD, Saunders PTK. Estrogen receptor related beta is expressed in human endometrium throughout the normal menstrual cycle. Hum Reprod. 2008;23(12):2782–2790. 70. Park S, Yoon S, Zhao Y, et al. Uterine development and fertility are dependent on gene dosage of the nuclear receptor coregulator REA. Endocrinology. 2012;153(8):3982–3994. 71. Vasquez YM, DeMayo FJ. Role of nuclear receptors in blastocyst implantation. Semin Cell Dev Biol. 2013;24(10–12):724–735. 72. Prossnitz ER, Barton M. The G-­protein-­coupled estrogen receptor GPER in health and disease.

73. Dennis MK, Burai R, Ramesh C, et al. In vivo effects of a GPR30 antagonist. Nat Chem Biol. 2009;5(6):421–427. 74. Dennis MK, Field AS, Burai R, et al. Identification of a GPER/ GPR30 antagonist with improved estrogen receptor counterselectivity. J Steroid Biochem Mol Biol. 2011;127(3–5):358–366. 75. Plante BJ, Lessey BA, Taylor RN, et al. G protein-­coupled estrogen receptor (GPER) expression in normal and abnormal endometrium. Reprod Sci. 2012;19(7):684–693. 76. Yuguchi H, Tanabe A, Hayashi A, et al. The expression status of G protein-­coupled receptor GPR30 is associated with the clinical characteristics of endometriosis. Endocr Res. 2013;38(4):223–231. 77. Mori T, Ito F, Matsushima H, et al. G protein-­ coupled estrogen receptor 1 agonist G-­ 1 induces cell cycle arrest in the mitotic phase, leading to apoptosis in endometriosis. Fertil Steril. 2015;103(5):1228–12235.e1. 78. Barrick CJ, Lenhart PM, Dackor RT, Nagle E, Caron KM. Loss of receptor activity-­modifying protein 3 exacerbates cardiac hypertrophy and transition to heart failure in a sex-­dependent manner. J Mol Cell Cardiol. 2012;52(1):165–174. 79. Liu T, Ogle TF. Signal transducer and activator of transcription 3 is expressed in the decidualized mesometrium of pregnancy and associates with the progesterone receptor through protein-­protein interactions. Biol Reprod. 2002;67(1):114–118. 80. Lydon JP, DeMayo FJ, Funk CR, et al. Mice lacking progesterone receptor exhibit pleiotropic reproductive abnormalities. Genes Dev. 1995;9(18):2266–2278. 81. Mote PA, Balleine RL, McGowan EM, Clarke CL. Heterogeneity of progesterone receptors A and B expression in human endometrial glands and stroma. Hum Reprod. 2000;15(suppl 3):48–56. 82. Boonyaratanakornkit V, Scott MP, Ribon V, et al. Progesterone receptor contains a proline-­rich motif that directly interacts with SH3 domains and activates c-­Src family tyrosine kinases. Mol Cell. 2001;8(2):269–280. 83. Boonyaratanakornkit V, Edwards DP. Receptor mechanisms of rapid extranuclear signalling initiated by steroid hormones. Essays Biochem. 2004;40:105–120. 84. Boonyaratanakornkit V, McGowan E, Sherman L, Mancini MA, Cheskis BJ, Edwards DP. The role of extranuclear signaling actions of progesterone receptor in mediating progesterone regulation of gene expression and the cell cycle. Mol Endocrinol. 2007;21(2):359–375. 85. Lessey BA, Yeh I, Castelbaum AJ, et al. Endometrial progesterone receptors and markers of uterine receptivity in the window of implantation. Fertil Steril. 1996;65(3):477–483. 86. Mote PA, Balleine RL, McGowan EM, Clarke CL. Colocalization of progesterone receptors A and B by dual immunofluorescent histochemistry in human endometrium during the menstrual cycle. J Clin Endocrinol Metab. 1999;84(8):2963–2971. 87. Vallejo G, La Greca AD, Tarifa-­Reischle IC, et al. CDC2 mediates progestin initiated endometrial stromal cell proliferation: a PR signaling to gene expression independently of its binding to chromatin. PLoS One. 2014;9(5):e97311. 88. Patel B, Elguero S, Thakore S, Dahoud W, Bedaiwy M, Mesiano S. Role of nuclear progesterone receptor isoforms in uterine pathophysiology. Hum Reprod Update. 2015;21(2):155–173. 89. Hirata S, Shoda T, Kato J, Hoshi K. Isoform/variant mRNAs for sex steroid hormone receptors in humans. Trends Endocrinol Metab. 2003;14(3):124–129. 90. Cork DM, Lennard TW, Tyson-­Capper AJ, Tyson-­Capper AJ. Alternative splicing and the progesterone receptor in breast cancer. Breast Cancer Res. 2008;10. 91. Wei LL, Hawkins P, Baker C, Norris B, Sheridan PL, Quinn PG. An amino-­terminal truncated progesterone receptor isoform, PRc, enhances progestin-­induced transcriptional activity. Mol Endocrinol. 1996;10(11):1379–1387. 92. Condon JC, Hardy DB, Kovaric K, Mendelson CR. Up-­regulation of the progesterone receptor (PR)-­C isoform in laboring myometrium by activation of nuclear factor-­kappaB may contribute to the onset of labor through inhibition of PR function. Mol Endocrinol. 2006;20(4):764–775. 93. Gellersen B, Fernandes MS, Brosens JJ. Non-­ genomic progesterone actions in female reproduction. Hum Reprod Update. 2009;15(1):119–138. 94. Cunha GR, Chung LW, Shannon JM, Reese BA. Stromal-­epithelial . 1980;22(1):19–42.

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract 95. Cunha GR, Cooke PS, Kurita T. Role of stromal-­epithelial interactions in hormonal responses. Arch Histol Cytol. 2004;67(5):417–434. 96. Cooke PS, Buchanan DL, Young P, et al. Stromal estrogen receptors mediate mitogenic effects of estradiol on uterine epithelium. Proc Natl Acad Sci U S A. 1997;94(12):6535–6540. 97. Conneely OM, Mulac-­ Jericevic B, Lydon JP. Progesterone-­ dependent regulation of female reproductive activity by two distinct progesterone receptor isoforms. Steroids. 2003;68(10–13):771–778. 98. Igarashi TM, Bruner-­Tran KL, Yeaman GR, et al. Reduced expression of progesterone receptor-­B in the endometrium of women with endometriosis and in cocultures of endometrial cells exposed to 2,3, 7,8-­tetrachlorodibenzo-­p-­dioxin. Fertil Steril. 2005;84(1):67–74. 99. Dressing GE, Goldberg JE, Charles NJ, Schwertfeger KL, Lange CA. Membrane progesterone receptor expression in mammalian tissues: a review of regulation and physiological implications. Steroids. 2011;76(1–2):11–17. 100. Zhu Y, Bond J, Thomas P. Identification, classification, and partial characterization of genes in humans and other vertebrates homologous to a fish membrane progestin receptor. Proc Natl Acad Sci U S A. 2003;100(5):2237–2242. 101. Karteris E, Zervou S, Pang Y, et al. Progesterone signaling in human myometrium through two novel membrane G protein-­ coupled receptors: potential role in functional progesterone withdrawal at term. Mol Endocrinol. 2006;20(7):1519–1534. 102. Cooke PS, Buchanan DL, Lubahn DB, Cunha GR. Mechanism of estrogen action: lessons from the estrogen receptor-­α knockout Mouse1. Biol Reprod. 1998;59(3):470–475. 103. Kurita T, Cooke PS, Cunha GR. Epithelial–stromal tissue interaction in paramesonephric (müllerian) epithelial differentiation. Dev Biol. 2001;240(1):194–211. 104. Pierro E, Minici F, Alesiani O, et al. Stromal-­epithelial interactions modulate estrogen responsiveness in normal human endometrium. Biol Reprod. 2001;64(3):831–838. 105. Dickson RB, Lippman ME. Estrogenic regulation of growth and polypeptide growth factor secretion in human breast carcinoma. Endocr Rev. 1987;8(1):29–43. 106. Hom YK, Young P, Wiesen JF, et al. Uterine and vaginal organ growth requires epidermal growth factor receptor signaling from stroma. Endocrinology. 1998;139(3):913–921. 107. Ritvos O, Ranta T, Jalkanen J, et al. Insulin-­like growth factor (IGF) binding protein from human decidua inhibits the binding and biological action of IGF-­I in cultured choriocarcinoma cells. Endocrinology. 1988;122(5):2150–2157. 108. Ghahary A, Murphy LJ. Uterine insulin-­like growth factor-­I receptors: regulation by estrogen and variation throughout the estrous cycle. Endocrinology. 1989;125(2):597–604. 109. Carlsson B, Billig H. Insulin-­like growth factor-­I gene expression during development and estrous cycle in the rat uterus. Mol Cell Endocrinol. 1991;77(1–3):175–180. 110. Rajkumar K, Krsek M, Dheen ST, Murphy LJ. Impaired glucose homeostasis in insulin-­like growth factor binding protein-­1 transgenic mice. J Clin Invest. 1996;98(8):1818–1825. 111. Sato T, Wang G, Hardy MP, Kurita T, Cunha GR, Cooke PS. Role of systemic and local IGF-­ I in the effects of estrogen on growth and epithelial proliferation of mouse uterus. Endocrinology. 2002;143(7):2673–2679. 112. Wetendorf M, DeMayo FJ. The progesterone receptor regulates implantation, decidualization, and glandular development via a complex paracrine signaling network. Mol Cell Endocrinol. 2012;357(1–2):108–118. 113. Kurita T, Young P, Brody JR, Lydon JP, O’Malley BW, Cunha GR. Stromal progesterone receptors mediate the inhibitory effects of progesterone on estrogen-­induced uterine epithelial cell deoxyribonucleic acid synthesis. Endocrinology. 1998;139(11):4708–4713. 114. Large MJ, DeMayo FJ. The regulation of embryo implantation and endometrial decidualization by progesterone receptor signaling. Mol Cell Endocrinol. 2012;358(2):155–165. 115. Lessey BA, Young SL. Homeostasis imbalance in the endometrium of women with implantation defects: the role of estrogen and progesterone. Semin Reprod Med. 2014;32(5):365–375. 116. Bell SC. Secretory endometrial and decidual proteins: studies and clinical significance of a maternally derived group of pregnancy-­ associated serum proteins. Hum Reprod. 1986;1(3):129–143. 117. Klinkova O, Hansen KA, Winterton E, Mark CJ, Eyster KM. Two-­ way communication between endometrial stromal cells and mono cytes. Reprod Sci

253.e3

118. Cunha GR, Bigsby RM, Cooke PS, Sugimura Y. Stromal-­epithelial interactions in adult organs. Cell Differ. 1985;17(3):137–148. 119. de Ziegler D, Bergeron C, Cornel C, et al. Effects of luteal estradiol on the secretory transformation of human endometrium and plasma gonadotropins. J Clin Endocrinol Metab. 1992;74(2):322–331. 120. de Ziegler D, Frydman R, Bouchard P. Contribution of estrogen to the morphology of “secretory” endometrium. Fertil Steril. 1995;63:1135. 121. Burney RO, Talbi S, Hamilton AE, et al. Gene expression analysis of endometrium reveals progesterone resistance and candidate susceptibility genes in women with endometriosis. Endocrinology. 2007;148(8):3814–3826. 122. Teague EMCO, Print CG, Hull ML. The role of microRNAs in endometriosis and associated reproductive conditions. Hum Reprod Update. 2010;16(2):142–165. 123. Teague O, Van Der Hoek EM, Van Der Hoek KH. MicroRNA-­ regulated pathways associated with endometriosis. Mol Endocrinol. 2009;23:265–275. 124. Burney RO, Hamilton AE, Aghajanova L, et al. MicroRNA expression profiling of eutopic secretory endometrium in women with versus without endometriosis. Mol Hum Reprod. 2009;15(10):625–631. 125. Joshi NR, Miyadahira EH, Afshar Y, et al. Progesterone resistance in endometriosis is modulated by the altered expression of microRNA-­ 29c and FKBP4. J Clin Endocrinol Metab. 2016. Published online jc.2016-­2076. 126. Rubel CA, Franco HL, Jeong JW, Lydon JP, DeMayo FJ. GATA2 is expressed at critical times in the mouse uterus during pregnancy. Gene Expr Patterns. 2012;12(5–6):196–203. 127. Li Q, Kannan A, DeMayo FJ, et al. The antiproliferative action of progesterone in uterine epithelium is mediated by Hand2. Science. 2011;331(6019):912–916. 128. Vasquez YM, Mazur EC, Li X, et al. FOXO1 is required for binding of PR on IRF4, novel transcriptional regulator of endometrial stromal decidualization. Mol Endocrinol. 2015;29(3):421–433. 129. Yang H, Zhou Y, Edelshain B, Schatz F, Lockwood CJ, Taylor HS. FKBP4 is regulated by HOXA10 during decidualization and in endometriosis. J Reprod Fertil. 2012;143(4):531–538. 130. Hewitt C, Goulding S, Eddy EH. Studies using the estrogen receptor alpha knockout uterus demonstrate that implantation but not decidualization-­ associated signaling is estrogen dependent. Biol Reprod. 2002;67:1268–1277. 131. Illera MJ, Cullinan E, Gui Y, Yuan L, Beyler SA, Lessey BA. Blockade of the alpha(v)beta(3) integrin adversely affects implantation in the mouse. Biol Reprod. 2000;62(5):1285–1290. 132. Illera MJ, Lorenzo PL, Gui YT, Beyler SA, Apparao KBC, Lessey BA. A role for alphavbeta3 integrin during implantation in the rabbit model. Biol Reprod. 2003;68(3):766–771. 133. Ahn JI, Yoo JY, Kim TH, Kim TH, et al. cAMP-­response element-­ binding 3-­like protein 1 (CREB3L1) is required for decidualization and its expression is decreased in women with endometriosis. Curr Mol Med. 2016;16:276–287. 134. Kim TH, Yoo JY, Wang Z, et al. ARID1A is essential for endometrial function during early pregnancy. PLoS Genet. 2015;11(9):e1005537. 135. Adesanya-­Famuyiwa OO, Zhou J, Wu G, Bondy C. Localization and sex steroid regulation of androgen receptor gene expression in rhesus monkey uterus. Obstet Gynecol. 1999;93(2):265–270. 136. Ito K, Suzuki T, Akahira JI, et al. Expression of androgen receptor and 5alpha-­reductases in the human normal endometrium and its disorders. Int J Cancer. 2002;99(5):652–657. 137. Apparao KBC, Lovely LP, Gui Y, Lininger RA, Lessey BA. Elevated endometrial androgen receptor expression in women with polycystic ovarian syndrome. Biol Reprod. 2002;66(2):297–304. 138. Cloke B, Christian M. The role of androgens and the androgen receptor in cycling endometrium. Mol Cell Endocrinol. 2012;358(2):166–175. 139. Aplin JD, Charlton AK, Ayad S. An immunohistochemical study of human endometrial extracellular matrix during the menstrual cycle and first trimester of pregnancy. Cell Tissue Res. 1988;253(1):231–240. 140. Iwahashi M, Muragaki Y, Ooshima A, Yamoto M, Nakano R. Alterations in distribution and composition of the extracellular matrix during decidualization of the human endometrium. J Reprod Fertil. 1996;108(1):147–155. 141. Brar AK, Handwerger S, Kessler CA, et al. Gene induction and catvitro human endometrial fibroPhysiol Genomics. 2001;7:135–148.

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253.e4 PART I  The Fundamentals of Reproduction 142. Schatz F, Guzeloglu-­Kayisli O, Arlier S, et al. The role of decidual cells in uterine hemostasis, menstruation, inflammation, adverse pregnancy outcomes and abnormal uterine bleeding. Hum Reprod Update. 2016;22:497–515. 143. Okulicz WC, Balsamo M. A double immunofluorescent method for simultaneous analysis of progesterone-­dependent changes in proliferation and the oestrogen receptor in endometrium of rhesus monkeys. J Reprod Fertil. 1993;99:545. 144. Okulicz WC, Ace CI, Scarrell R. Zonal changes in proliferation in the rhesus endometrium during the late secretory phase and menses. Proc Soc Exp Biol Med. 1997;214:132. 145. Slayden OD, Brenner RM. Hormonal regulation and localization of estrogen, progestin and androgen receptors in the endometrium of nonhuman primates: effects of progesterone receptor antagonists. Arch Histol Cytol. 2004;67:393–409. 146. Gargett CE, Chan RW, Schwab KE. Endometrial stem cells. Curr Opin Obstet Gynecol. 2007;19:377–383. 147. Tulac S, Nayak NR, Kao LC, et al. Identification, characterization, and regulation of the canonical WNT signaling pathway in human endometrium. J Clin Endocrinol Metab. 2003;88:3860–3866. 148. Tulac S, Overgaard MT, Hamilton AE, et al. Dickkopf-­1, an inhibitor of WNT signaling, is regulated by progesterone in human endometrial stromal cells. J Clin Endocrinol Metab. 2006;91:1453–1461. 149. Kao LC, Germeyer A, Tulac S, et al. Expression profiling of endometrium from women with endometriosis reveals candidate genes for disease-­based implantation failure and infertility. Endocrinology. 2003;144:2870–2881. 150. Slayden OD, Keator CS. Role of progesterone in nonhuman primate implantation. Semin Reprod Med. 2007;25:418–430. 151. Wynn RM. Histology and ultrastructure of the human endometrium. In: Wynn RM, Jolie WP, eds. Biology of the Uterus. New York: Plenum Publishing; 1977:341–376. 152. Isaac C, Pollard JW, Meier UT. Intranuclear endoplasmic reticulum induced by Nopp140 mimics the nucleolar channel system of human endometrium. J Cell Sci. 2001;114:4253–4264. 153. King A. Uterine leukocytes and decidualization. Hum Reprod Update. 2000;6:28–36. 154. Chan RW, Gargett CE. Identification of label-­retaining cells in mouse endometrium. Stem Cell. 2006;24:1529–1538. 155. Cervello I, Gil-­Sanchis C, Mas A, et al. Human endometrial side population cells exhibit genotypic, phenotypic and functional features of somatic stem cells. PLoS One. 2010;5:e10964. 156. Salamonsen LA, Lathbury LJ. Endometrial leukocytes and menstruation. Hum Reprod Update. 2000;6:16–27. 157. Evans J, Salamonsen LA. Inflammation, leukocytes and menstruation. Rev Endocr Metab Disord. 2012;13:277–288. 158. Curry TE, Osteen KG. The matrix metalloproteinase system: changes, regulation, and impact throughout the ovarian and uterine reproductive cycle. Endocr Rev. 2003;24:428–465. 159. Markee JE. Menstruation in intraocular endometrial transplants in the Rhesus monkey. Am J Obstet Gynecol. 1978;131:558–559. 160. Fraser IS, McCarron G, Hutton B, et al. Endometrial blood flow measured by xenon 133 clearance in women with normal menstrual cycles and dysfunctional uterine bleeding. Am J Obstet Gynecol. 1987;156:158–166. 161. Zhang J, Salamonsen LA. Expression of hypoxia-­inducible factors in human endometrium and suppression of matrix metalloproteinases under hypoxic conditions do not support a major role for hypoxia in regulating tissue breakdown at menstruation. Hum Reprod. 2002;17:265–274. 162. Marbaix E, Kokorine I, Moulin P, et al. Menstrual breakdown of human endometrium can be mimicked in vitro and is selectively and reversibly blocked by inhibitors of matrix metalloproteinases. Proc Natl Acad Sci U S A. 1996;93:9120–9125. 163. Rodgers WH, Matrisian LM, Giudice LC, et al. Patterns of matrix metalloproteinase expression in cycling endometrium imply differential functions and regulation by steroid hormones. J Clin Invest. 1994;94:946–953. 164. Irwin JC, Kirk D, Gwatkin RB, et al. Human endometrial matrix metalloproteinase-­2, a putative menstrual proteinase—Hormonal regulation in cultured stromal cells and messenger RNA expression during the menstrual cycle. J Clin Invest. 1996;97:438–447. 165. Henriet P, Cornet PB, Lemoine P, et al. Circulating ovarian steroids and endometrial matrix metalloproteinases (MMPs). Ann N Y Acad Sci. 2002;955:119–138. discussion 157–158, 396–406.

166. Otsuki Y. Apoptosis in human endometrium: apoptotic detection methods and signaling. Med Electron Microsc. 2001;34:166–173. 167. Kokawa K, Shikone T, Nakano R. Apoptosis in the human uterine endometrium during the menstrual cycle. J Clin Endocrinol Metab. 1996;81:4144–4147. 168. Konno R, Yamakawa H, Utsunomiya H, et al. Expression of survivin and Bcl-­ 2 in the normal human endometrium. Mol Hum Reprod. 2000;6:529–534. 169. Lessey BA, Lovely LP, McAdams D, et al. Randomized prospective comparison of hCG versus progesterone to prevent apoptosis in late luteal phase endometrium. J Clin Endocrinol Metab. 2000 (in preparation). 170. Ueda M, Yamashita Y, Takehara M, et al. Survivin gene expression in endometriosis. J Clin Endocrinol Metab. 2002;87:3452–3459. 171. Maybin JA, Critchley HO. Menstrual physiology: implications for endometrial pathology and beyond. Hum Reprod Update. 2015;21:748–761. 172. Zhang J, Hampton AL, Nie G, et al. Progesterone inhibits activation of latent matrix metalloproteinase (MMP)-­2 by membrane-­type 1 MMP: enzymes coordinately expressed in human endometrium. Biol Reprod. 2000;62:85–94. 173. Salamonsen LA, Woolley DE. Menstruation: induction by matrix metalloproteinases and inflammatory cells. J Reprod Immunol. 1999;44:1–27. 174. Hutchinson JL, Rajagopal SP, Sales KJ, et al. Molecular regulators of resolution of inflammation: potential therapeutic targets in the reproductive system. Reproduction. 2011;142:15–28. 175. Macdonald LJ, Boddy SC, Denison FC, et al. A role for lipoxin A(4) as an anti-­inflammatory mediator in the human endometrium. Reproduction. 2011;142:345–352. 176. Bannenberg G, Serhan CN. Specialized pro-­resolving lipid mediators in the inflammatory response: an update. Biochim Biophys Acta. 2010;1801:1260–1273. 177. Russell R, Gori I, Pellegrini C, et al. Lipoxin A4 is a novel estrogen receptor modulator. FASEB J. 2011;25:4326–4337. 178. Taylor HS. Endometrial cells derived from donor stem cells in bone marrow transplant recipients. JAMA. 2004;292:81–85. 179. Gargett CE, Schwab KE, Brosens JJ, et al. Potential role of endometrial stem/progenitor cells in the pathogenesis of early-­onset endometriosis. Mol Hum Reprod. 2014;20:591–598. 180. Santamaria X, Massasa EE, Feng Y, et al. Derivation of insulin producing cells from human endometrial stromal stem cells and use in the treatment of murine diabetes. Mol Ther. 2011;19:2065–2071. 181. Wolff EF, Gao XB, Yao KV, et al. Endometrial stem cell transplantation restores dopamine production in a Parkinson’s disease model. J Cell Mol Med. 2011;15:747–755. 182. Masuda H, Anwar SS, Buhring HJ, et al. A novel marker of human endometrial mesenchymal stem-­like cells. Cell Transplant. 2012;21(10):2201–2214. 183. Cervello I, Mas A, Gil-­Sanchis C, et al. Reconstruction of endometrium from human endometrial side population cell lines. PLoS One. 2011;6:e21221. 184. Chan RW, Kaitu’u-­Lino T, Gargett CE. Role of label-­retaining cells in estrogen-­ induced endometrial regeneration. Reprod Sci. 2012;19:102–114. 185. Gargett CE, Healy DL. Generating receptive endometrium in Asherman’s syndrome. J Hum Reprod Sci. 2011;4:49–52. 186. Jang HY, Myoung SM, Choe JM, et al. Effects of autologous platelet-­rich plasma on regeneration of damaged endometrium in female rats. Yonsei Med J. 2017;58(6):1195. 187. Kim JH, Park M, Paek JY, Lee WS, Song H, Lyu SW. Intrauterine infusion of human platelet-­rich plasma improves endometrial regeneration and pregnancy outcomes in a Murine model of Asherman’s syndrome. Front Physiol. 2020;11:105. 188. de Miguel-­Gómez L, López-­ Martínez S, Campo H, et al. Comparison of different sources of platelet-­rich plasma as treatment option for infertility-­causing endometrial pathologies. Fertil Steril. 2021;115(2):490–500. 189. Zadehmodarres S, Salehpour S, Saharkhiz N, Nazari L. Treatment of thin endometrium with autologous platelet-­rich plasma: a pilot study. JBRA Assist Reprod. 2017;21(1). 190. Kim H, Shin JE, Koo HS, Kwon H, Choi DH, Kim JH. Effect of autologous platelet-­rich plasma treatment on refractory thin endometrium during the frozen embryo transfer cycle: a pilot study.

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract 191. Cervello I, Martinez-­ Conejero JA, Horcajadas JA, et al. Identification, characterization and co-­localization of label-­retaining cell population in mouse endometrium with typical undifferentiated markers. Hum Reprod. 2007;22:45–51. 192. Cervello I, Gil-­Sanchis C, Mas A, et al. Bone marrow-­derived cells from male donors do not contribute to the endometrial side population of the recipient. PLoS One. 2012;7:e30260. 193. Schwab KE, Gargett CE. Co-­expression of two perivascular cell markers isolates mesenchymal stem-­like cells from human endometrium. Hum Reprod. 2007;22:2903–2911. 194. Haber DA. Telomeres, cancer, and immortality. N Engl J Med. 1995;332:955–956. 195. Kyo S, Takakura M, Kohama T, et al. Telomerase activity in human endometrium. Cancer Res. 1997;57:610–614. 196. Yokoyama Y, Takahashi Y, Morishita S, et al. Telomerase activity in the human endometrium throughout the menstrual cycle. Mol Hum Reprod. 1998;4:173–177. 197. Williams CD, Boggess JF, LaMarque LR, et al. A prospective, randomized study of endometrial telomerase during the menstrual cycle. J Clin Endocrinol Metab. 2001;86:3912–3917. 198. Yokoyama Y, Takahashi Y, Shinohara A, et al. Telomerase activity is found in the epithelial cells but not in the stromal cells in human endometrial cell culture. Mol Hum Reprod. 1998;4:985–989. 199. Gargett CE, Ye L. Endometrial reconstruction from stem cells. Fertil Steril. 2012;98:11–20. 200. Hoffmeyer K, Raggioli A, Rudloff S, et al. WNT/beta-­catenin signaling regulates telomerase in stem cells and cancer cells. Science. 2012;336:1549–1554. 201. Hapangama DK, Turner MA, Drury J, et al. Aberrant expression of regulators of cell-­fate found in eutopic endometrium is found in matched ectopic endometrium among women and in a baboon model of endometriosis. Hum Reprod. 2010;25:2840–2850. 202. Gargett CE, Rogers PA. Human endometrial angiogenesis. Reproduction. 2001;121:181–186. 203. Smith SK. Angiogenesis, vascular endothelial growth factor and the endometrium. Hum Reprod Update. 1998;4:509–519. 204. Krikun G, Schatz F, Lockwood CJ. Endometrial angiogenesis: from physiology to pathology. Ann N Y Acad Sci. 2004;1034:27–35. 205. Gambino LS, Wreford NG, Bertram JF, et al. Angiogenesis occurs by vessel elongation in proliferative phase human endometrium. Hum Reprod. 2002;17:1199–1206. 206. Koga K, Osuga Y, Tsutsumi O, et al. Demonstration of angiogenin in human endometrium and its enhanced expression in endometrial tissues in the secretory phase and the decidua. J Clin Endocrinol Metab. 2001;86:5609–5614. 207. Ancelin M, Buteau-­Lozano H, Meduri G, et al. A dynamic shift of VEGF isoforms with a transient and selective progesterone-­induced expression of VEGF189 regulates angiogenesis and vascular permeability in human uterus. Proc Natl Acad Sci U S A. 2002;99:6023–6028. 208. Moller B, Rasmussen C, Lindblom B, et al. Expression of the angiogenic growth factors VEGF, FGF-­2, EGF and their receptors in normal human endometrium during the menstrual cycle. Mol Hum Reprod. 2001;7:65–72. 209. Sugino N, Kashida S, Takiguchi S, et al. Expression of vascular endothelial growth factor and its receptors in the human corpus luteum during the menstrual cycle and in early pregnancy. J Clin Endocrinol Metab. 2000;85:3919–3924. 210. Krussel JS, Casan EM, Raga R, et al. Expression of mRNA for vascular endothelial growth factor transmembraneous receptors Flt1 and KDR, and the soluble receptor sflt in cycling human endometrium. Mol Hum Reprod. 2001;5:452–458. 211. Meduri G, Bausero P, Perrot-­ Applanat M. Expression of vascular endothelial growth factor receptors in the human endometrium: modulation during the menstrual cycle. Biol Reprod. 2000;62:439–447. 212. Molitoris KH, Kazi AA, Koos RD. Inhibition of oxygen-­induced hypoxia-­ inducible factor-­ 1alpha degradation unmasks estradiol induction of vascular endothelial growth factor expression in ECC-­1 cancer cells in vitro. Endocrinology. 2009;150:5405–5414. 213. Koos RD. Minireview: putting physiology back into estrogens’ mechanism of action. Endocrinology. 2011;152:4481–4488. 214. Critchley HO, Osei J, Henderson TA, et al. Hypoxia-­inducible factor-­1alpha expression in human endometrium and its regulation by prostaglandin E-­series prostanoid receptor 2 (EP2). Endocrinology. 2006;147:744–753.

253.e5

215. Graubert MD, Ortega MA, Kessel B, et al. Vascular repair after menstruation involves regulation of vascular endothelial growth factor-­receptor phosphorylation by sFLT-­1. Am J Pathol. 2001;158:1399–1410. 216. Brooks PC, Clark RAF, Cheresh DA. Requirement of vascular integrin alpha v/beta 3 for angiogenesis. Science. 1994;264:569–571. 217. Hewett P, Nijjar S, Shams M, et al. Down-­regulation of angiopoietin-­1 expression in menorrhagia. Am J Pathol. 2002;160:773–780. 218. Gannon BJ, Verco CJ, Carati CJ. Human endometrial perfusion after tubal occlusion. Hum Reprod. 1998;13:445–449. 219. Casslen B, Andersson A, Nilsson IM, et al. Hormonal regulation of the release of plasminogen activators and of a specific activator inhibitor from endometrial tissue in culture. Proc Soc Exp Biol Med. 1986;182:419–424. 220. Rudolph-­Owen LA, Slayden OD, Matrisian LM, et al. Matrix metalloproteinase expression in Macaca mulatta endometrium: evidence for zone-­specific regulatory tissue gradients. Biol Reprod. 1998;59:1349–1359. 221. Tabibzadeh S. Role of EBAF/Lefty in Implantation and Uterine Bleeding. Ernst Schering Res Found Workshop; 2005:159–189. 222. Kothapalli R, Buyuksal I, Wu SQ, et al. Detection of ebaf, a novel human gene of the transforming growth factor beta superfamily— Association of gene expression with endometrial bleeding. J Clin Invest. 1997;99:2342–2350. 223. Tabibzadeh S, Lessey B, Satyaswaroop PG. Temporal and site-­ specific expression of transforming growth factor-­beta4 in human endometrium. Mol Hum Reprod. 1998;4:595–602. 224. Tabibzadeh S, Mason JM, Shea W, et al. Dysregulated expression of ebaf, a novel molecular defect in the endometria of patients with infertility. J Clin Endocrinol Metab. 2000;85:2526–2536. 225. Cornet PB, Picquet C, Lemoine P, et al. Regulation and function of LEFTY-­A/EBAF in the human endometrium. mRNA expression during the menstrual cycle, control by progesterone, and effect on matrix metalloprotineases. J Biol Chem. 2002;277:42496–42504. 226. Jokimaa V, Oksjoki S, Kujari H, et al. Expression patterns of cathepsins B, H, K, L and S in the human endometrium. Mol Hum Reprod. 2001;7:73–78. 227. Akerlund A. Vascularization of human endometrium: uterine blood flow in healthy condition and in primary dysmenorrhoea. Ann N Y Acad Sci. 1994;734:47–55. 228. Bieglmayer C, Hofer G, Kainz C, et al. Concentrations of various arachidonic acid metabolites in menstrual fluid are associated with menstrual pain and are influenced by hormonal contraceptives. Gynecol Endocrinol. 1995;9:307–312. 229. Casey ML, Hemsell DL, MacDonald PC, et al. NAD+-­dependent 15-­hydroxyprostaglandin dehydrogenase activity in human endometrium. Prostaglandins. 1980;19:115–122. 230. Maybin JA, Hirani N, Jabbour HN, et al. Novel roles for hypoxia and prostaglandin E2 in the regulation of IL-­8 during endometrial repair. Am J Pathol. 2011;178:1245–1256. 231. Schatz F, Aigner S, Papp C, et al. Plasminogen activator activity during decidualization of human endometrial stromal cells is regulated by plasminogen activator inhibitor 1. J Clin Endocrinol Metab. 1995;80:2504–2510. 232. Lockwood CJ. Mechanisms of normal and abnormal endometrial bleeding. Menopause. 2011;18:408–411. 233. Lockwood CJ. Regulation of plasminogen activator inhibitor 1 expression by interaction of epidermal growth factor with progestin during decidualization of human endometrial stromal cells. Am J Obstet Gynecol. 2001;184:798–804. Discussion 804–805. 234. Papp C, Schatz F, Krikun G, et al. Biological mechanisms underlying the clinical effects of mifepristone (RU 486) on the endometrium. Early Pregnancy. 2000;4:230–239. 235. Salamonsen LA, Edgell T, Rombauts LJ, et al. Proteomics of the human endometrium and uterine fluid: a pathway to biomarker discovery. Fertil Steril. 2013;99:1086–1092. 236. Scotchie JG, Fritz MA, Mocanu M, et al. Proteomic analysis of the luteal endometrial secretome. Reprod Sci. 2009;16:883–893. 237. Salamonsen LA, Evans J, Nguyen HP, et al. The microenvironment of human implantation: determinant of reproductive success. Am J Reprod Immunol. 2016;75:218–225. 238. Paria BC, Reese J, Das SK, et al. Deciphering the cross-­talk of implantation: advances and challenges. Science. 2002;296:2185–2188. 239. Paria BC, Song H, Dey SK. Implantation: molecular basis of . 2001;45:597–605.

10

253.e6 PART I  The Fundamentals of Reproduction 240. Bhurke AS, Bagchi IC, Bagchi MK. Progesterone-­regulated endometrial factors controlling implantation. Am. J Reprod Immunol. 2016;75:237–245. 241. Burton GJ, Watson AL, Hempstock J, et al. Uterine glands provide histiotrophic nutrition for the human fetus during the first trimester of pregnancy. J Clin Endocrinol Metab. 2002;87:2954–2959. 242. Langmann T, Mauerer R, Zahn A, et al. Real-­ time reverse transcription-­ PCR expression profiling of the complete human ATP-­ binding cassette transporter superfamily in various tissues. Clin Chem. 2003;49:230–238. 243. Spencer TE. Biological roles of uterine glands in pregnancy. Semin Reprod Med. 2014;32(5):346–357. 244. Rogers PAW, Polson D, Murphy CR, Hosie M, Susil B, Leoni M. Correlation of endometrial histology, morphometry, and ultrasound appearance after different stimulation protocols for in vitro fertilization. Fertil Steril. 1991;55(3):583–587. 245. Burton GJ, Watson AL, Hempstock J, Skepper JN, Jauniaux E. Uterine glands provide histiotrophic nutrition for the human fetus during the first trimester of pregnancy. J Clin Endocrinol Metab. 2002;87(6):2954–2959. 246. Seppäla M, Suikkari AM, Julkunen M. Human endometrial proteins. Reprod Nutr Dev. 1988;28:1649–1654. 247. Bell SC, Patel SR, Kirwan PH, et al. Protein synthesis and secretion by the human endometrium during the menstrual cycle and the effect of progesterone in vitro. J Reprod Fertil. 1986;77:221–231. 248. Seppäla M, Koistinen R, Rutanen EM. Uterine endocrinology and paracrinology: insulin-­like growth factor binding protein-­1 and placental protein 14 revisited. Hum Reprod. 1994;9:917–925. 249. Seppala M, Taylor RN, Koistinen H, et al. Glycodelin: a major lipocalin protein of the reproductive axis with diverse actions in cell recognition and differentiation. Endocr Rev. 2002;23:401–430. 250. Dell A, Morris HR, Easton RL, et al. Structural analysis of the oligosaccharides derived from glycodelin, a human glycoprotein with potent immunosuppressive and contraceptive activities. J Biol Chem. 1995;270:24116–24126. 251. Oehninger S, Coddington CC, Hodgen GD, et al. Factors affecting fertilization: endometrial placental protein 14 reduces the capacity of human spermatozoa to bind to the human zona pellucida. Fertil Steril. 1995;63:377–383. 252. Lam KK, Chiu PC, Chung MK, et al. Glycodelin-­A as a modulator of trophoblast invasion. Hum Reprod. 2009;24:2093–2103. 253. Stewart DR, Erikson MS, Erikson ME, et al. The role of relaxin in glycodelin secretion. J Clin Endocrinol Metab. 1997;82:839–846. 254. Klentzeris LD, Bulmer JN, Seppala M, et al. Placental protein 14 in cycles with normal and retarded endometrial differentiation. Hum Reprod. 1994;9:394–398. 255. McKenna NJ, O’Malley BW. Combinatorial control of gene expression by nuclear receptors and coregulators. Cell. 2002;108:465–474. 256. Takai N, Desmond JC, Kumagai T, et al. Histone deacetylase inhibitors have a profound antigrowth activity in endometrial cancer cells. Clin Cancer Res. 2004;10:1141–1149. 257. Uchida H, Maruyama T, Ohta K, et al. Histone deacetylase inhibitor-­induced glycodelin enhances the initial step of implantation. Hum Reprod. 2007;22:2615–2622. 258. Jakubowicz DJ, Essah PA, Seppala M, et al. Reduced serum glycodelin and insulin-­like growth factor-­binding protein-­1 in women with polycystic ovary syndrome during first trimester of pregnancy. J Clin Endocrinol Metab. 2004;89:833–839. 259. Raymond EG, Lovely LP, Chen-­Mok M, et al. Effect of the Yuzpe regimen of emergency contraception on markers of endometrial receptivity. Hum Reprod. 2000;15:2351–2355. 260. Mandelin E, Koistinen H, Koistinen R, et al. Levonorgestrel-­ releasing intrauterine device-­wearing women express contraceptive glycodelin A in endometrium during midcycle: another contraceptive mechanism? Hum Reprod. 1997;12:2671. 261. Giudice LC. Growth factors and growth modulators in human uterine endometrium: their potential relevance to reproductive medicine. Fertil Steril. 1994;61:1–17. 262. Crossey PA, Pillai CC, Miell JP. Altered placental development and intrauterine growth restriction in IGF binding protein-­1 transgenic mice. J Clin Invest. 2002;110:411–418. 263. Gleeson LM, Chakraborty C, McKinnon T, et al. Insulin-­like growth factor-­ binding protein 1 stimulates human trophoblast migration by signaling through alpha 5 beta 1 integrin via mitogen-­ activated protein Kinase pathway. Metab. 2001;86:2484–2493.

264. Thraikill KM, Clemmons DR, Busby Jr WH, et al. Differential regulation of insulin-­like growth factor binding protein secretion from human decidual cells by IGF-­I, insulin, and relaxin. J Clin Invest. 1990;86:878–883. 265. Gao J, Mazella J, Tseng L. Activation of the human IGFBP-­1 gene promoter by progestin and relaxin in primary culture of human endometrial stromal cells. Mol Cell Biol. 1994;104:29–46. 266. Tazuke SI, Mazure NM, Sugawara J, et al. Hypoxia stimulates insulin-­like growth factor binding protein 1 (IGFBP-­1) gene expression in HepG2 cells: a possible model for IGFBP-­1 expression in fetal hypoxia. Proc Natl Acad Sci U S A. 1998;95:10188–10193. 267. Kim JJ, Taylor HS, Akbas GE, et al. Regulation of insulin-­like growth factor binding protein-­1 promoter activity by FKHR and HOXA10 in primate endometrial cells. Biol Reprod. 2003;68:24–30. 268. Licht P, Russu V, Lehmeyer S, et al. Intrauterine microdialysis reveals cycle-­ dependent regulation of endometrial insulin-­ like growth factor binding protein-­ 1 secretion by human chorionic gonadotropin. Fertil Steril. 2002;78:252–258. 269. Savaris RF, Hamilton AE, Lessey BA, et al. Endometrial gene expression in early pregnancy: lessons from human ectopic pregnancy. Reprod Sci. 2008;15:797–816. 270. Kim JJ, Taylor HS, Lu Z, et al. Altered expression of HOXA10 in endometriosis: potential role in decidualization. Mol Hum Reprod. 2007;13:323–332. 271. Jain A, Karadag A, Fohr B, et al. Three SIBLINGs (small integrin-­ binding ligand, N-­linked glycoproteins) enhance factor H’s cofactor activity enabling MCP-­like cellular evasion of complement-­mediated attack. J Biol Chem. 2002;277:13700–13708. 272. Fisher LW, Torchia DA, Fohr B, et al. Flexible structures of SIBLING proteins, bone sialoprotein, and osteopontin. Biochem Biophys Res Commun. 2001;280:460–465. 273. Apparao KB, Murray MJ, Fritz MA, et al. Osteopontin and its receptor alphavbeta(3) integrin are coexpressed in the human endometrium during the menstrual cycle but regulated differentially. J Clin Endocrinol Metab. 2001;86:4991–5000. 274. Johnson GA, Spencer TE, Burghardt RC, et al. Progesterone modulation of osteopontin gene expression in the ovine uterus. Biol Reprod. 2000;62:1315–1321. 275. Fedarko NS, Fohr B, Robey PG, et al. Factor H binding to bone sialoprotein and osteopontin enables tumor cell evasion of complement-­mediated attack. J Biol Chem. 2000;275:16666–16672. 276. Maslar IA, Ansbacher R. Effects of progesterone on decidual prolactin production by organ cultures of human endometrium. Endocrinology. 1986;118:2102. 277. Kauma S, Shapiro SS. Immunoperoxidase localization of prolactin in endometrium during normal menstrual, luteal phase defect, and corrected luteal phase defect cycles. Fertil Steril. 1986;46:37–41. 278. Ohata Y, Harada T, Fujii A, et al. Menstrual cycle-­specific inhibition of endometrial stromal cell proliferation by oncostatin M. Mol Hum Reprod. 2001;7:665–670. 279. Tennekoon KH, Eswaramohan T, Karunanayake EH. Effect of leptin on prolactin and insulin-­like growth factor-­I secretion by cultured rat endometrial stromal cells. Fertil Steril. 2007;88:193–199. 280. Eyal O, Jomain JB, Kessler C, et al. Autocrine prolactin inhibits human uterine decidualization: a novel role for prolactin. Biol Reprod. 2007;76:777–783. 281. Tawadros N, Salamonsen LA, Dimitriadis E, et al. Facilitation of decidualization by locally produced ghrelin in the human endometrium. Mol Hum Reprod. 2007;13:483–489. 282. Kutsukake M, Ishihara R, Yoshie M, et al. Involvement of insulin-­ like growth factor-­ binding protein-­ related protein 1 in decidualization of human endometrial stromal cells. Mol Hum Reprod. 2007;13:737–743. 283. Takemura T, Murata Y, Hino S, et al. Heparin-­binding EGF-­ like growth factor is expressed by mesangial cells and is involved in mesangial proliferation in glomerulonephritis. J Pathol. 1999;189:431–438. 284. Reese J, Binart N, Brown N, et al. Implantation and decidualization defects in prolactin receptor (PRLR)-­deficient mice are mediated by ovarian but not uterine PRLR. Endocrinology. 2000;141:1872–1881. 285. Jabbour HN, Gubbay O, Critchley HO. Prolactin action and signaling in the human endometrium. Reprod Med Rev. 2002;10:117–132. 286. Carson DD, Bagchi I, Dey SK, et al. Embryo implantation. Dev Biol. 2000;223:217–237. J Reprod Fertil.

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract 288. Psychoyos A. Hormonal control of ovoimplantation. Vitam Horm. 1973;31:201–256. 289. Psychoyos A. Uterine receptivity for nidation. Ann N Y Acad Sci. 1986;476:36–42. 290. Pope WF. Uterine asynchrony: a cause of embryonic loss. Biol Reprod. 1988;39:999–1003. 291. Hodgen GD. Surrogate embryo transfer combined with estrogen-­ progesterone therapy in monkeys: implantation, gestation, and delivery without ovaries. JAMA. 1983;250:2167–2171. 292. Navot D, Bergh P. Preparation of the human endometrium for implantation. Ann N Y Acad Sci. 1991;622:212–219. 293. Wilcox AJ, Baird DD, Wenberg CR. Time of implantation of the conceptus and loss of pregnancy. N Engl J Med. 1999;340:1796–1799. 294. Cha J, Sun X, Dey SK. Mechanisms of implantation: strategies for successful pregnancy. Nat Med. 2012;18:1754–1767. 295. Rubel CA, Wu SP, Lin L, et al. A Gata2-­dependent transcription network regulates uterine progesterone responsiveness and endometrial function. Cell Rep. 2016;17:1414–1425. 296. Lessey BA. Assessment of endometrial receptivity. Fertil Steril. 2011;96(3):522–529. 297. Achache H, Revel A. Endometrial receptivity markers, the journey to successful embryo implantation. Hum Reprod Update. 2006;12(6):731–746. 298. Díaz-­Gimeno P, Horcajadas JA, Martínez-­ Conejero JA, et al. A genomic diagnostic tool for human endometrial receptivity based on the transcriptomic signature. Fertil Steril. 2011;95(1):50–60. 60.e1–e15. 299. Ruiz-­Alonso M, Blesa D, Díaz-­Gimeno P, et al. The endometrial receptivity array for diagnosis and personalized embryo transfer as a treatment for patients with repeated implantation failure. Fertil Steril. 2013;100(3):818–824. 300. Zhou Y, Fisher SJ, Janatpour M, et al. Human cytotrophoblasts adopt a vascular phenotype as they differentiate—A strategy for successful endovascular invasion? J Clin Invest. 1997;99:2139–2151. 301. Hertig AT, Rock J, Adams EC. A description of 34 human ova within the first 17 days of development. Am J Anat. 1956;98:435–493. 302. Mazur MT, Duncan DA, Younger JB. Endometrial biopsy in the cycle of conception: histologic and lectin histochemical evaluation. Fertil Steril. 1989;51:764–769. 303. Thrasher TV, Richart RM. Ultrastructure of the Arias-­Stella reaction. Am J Obstet Gynecol. 1972;112:113–120. 304. Arias-­Stella J. The Arias-­ Stella reaction: facts and fancies four decades after. Adv Anat Pathol. 2002;9:12–23. 305. Burton GJ, Jauniaux E, Charnock-­Jones DS. Human early placental development: potential roles of the endometrial glands. Placenta. 2007;28(suppl A):S64–S69. 306. Tan J, Raja S, Davis MK, Tawfik O, Dey SK, Das SK. Evidence for coordinated interaction of cyclin D3 with p21 and cdk6 in directing the development of uterine stromal cell decidualization and polyploidy during implantation. Mech Dev. 2002;111(1–2):99–113. 307. Dey SK, Lim H, Das SK, et al. Molecular cues to implantation. Endocr Rev. 2004;25(3):341–373. 308. Gellersen B, Brosens IA, Brosens JJ. Decidualization of the human endometrium: mechanisms, functions, and clinical perspectives. Semin Reprod Med. 2007;25(6):445–453. 309. Das SK. Regional development of uterine decidualization: molecular signaling by Hoxa-­10: HOXA-­10 signaling in decidualization. Mol Reprod Dev. 2010;77(5):387–396. 310. Qi QR, Zhao XY, Zuo RJ, et al. Involvement of atypical transcription factor E2F8 in the polyploidization during mouse and human decidualization. Cell Cycle. 2015;14(12):1842–1858. 311. Buzzio OL, Lu Z, Miller CD, et al. FOXO1A differentially regulates genes of decidualization. Endocrinology. 2006;147:3870–3876. 312. Gaytan F, Morales C, Barreiro ML, et al. Expression of growth hormone secretagogue receptor type 1a, the functional ghrelin receptor, in human ovarian surface epithelium, mullerian duct derivatives, and ovarian tumors. J Clin Endocrinol Metab. 2005;90:1798–1804. 313. Aghajanova L, Rumman A, Altmae S, et al. Diminished endometrial expression of ghrelin and ghrelin receptor contributes to infertility. Reprod Sci. 2010;17:823–832. 314. Enders AC, Welsh AO. Structural interactions of trophoblast and uterus during hemochorial placenta formation. J Exp Zool. 1993;266:578–587. 315. Larsen JF, Hubinot PO, Leroy F, et al. Electron microscopy of nidation in the rabbit and observations on the human trophoblastic invasion. In: Karger S and Prolactin. Basel; 1970:38–51.

253.e7

316.  Early development of the human placenta. In: Bernirschke K, Kaufmann P, Bernirschke K, et al., eds. Pathology of the Human Placenta. Springer-­Verlag; 1991:13–21. 317.  Receptor concept in implantation research. In: Yoshinaga K, Yoshinaga K, Mori T, eds. Development of Preimplantation Embryos and Their Environment. Alan Liss, Inc; 1989:379–387. 318. Lessey BA, Damjanovich L, Coutifaris C, et al. Integrin adhesion molecules in the human endometrium. Correlation with the normal and abnormal menstrual cycle. J Clin Invest. 1992;90:188–195. 319. Fukuda MN, Sato T, Nakayama J, et al. Trophinin and tastin, a novel cell adhesion molecule complex with potential involvement in embryo implantation. Genes Dev. 1995;9:1199–1210. 320. Chobotova K, Spyropoulou I, Carver J, et al. Heparin-­binding epidermal growth factor and its receptor ErbB4 mediate implantation of the human blastocyst. Mech Dev. 2002;119:137–144. 321. Genbacev OD, Prakobphol A, Foulk RA, et al. Trophoblast L-­selectin-­mediated adhesion at the maternal-­fetal interface. Science. 2003;299:405–408. 322. Carson DD, Julian J, Lessey BA, et al. MUC1 is a scaffold for selectin ligands in the human uterus. Front Biosci. 2006;11:2903–2908. 323. Anderson TL, Hoffman LH. Alerations in epithelial glycocalyx of rabbit uteri during early pseudopregnancy and pregnancy and following ovariectomy. Am J Anat. 1984;171:321–334. 324. Peverini S, Kramer B. Superovulation affects glucosamine trisaccharides in the glycocalyx of the rat endometrium at the time of implantation. J Anat. 1995;187(Pt 2):487–490. 325. Aplin JD. MUC-­1 glycosylation in endometrium: possible roles of the apical glycocalyx at implantation. Hum Reprod. 1999;14(suppl 2):17–25. 326. DeSouza MM, Surveyor GA, Price RE, et al. MUC1/episialin: a critical barrier in the female reproductive tract. J Reprod Immunol. 1999;45:127–158. 327. Achache H, Revel A. Endometrial receptivity markers, the journey to successful embryo implantation. Hum Reprod Update. 2006;12:731–746. 328. Donaghay M, Lessey BA. Uterine receptivity: alterations associated with benign gynecological disease. Semin Reprod Med. 2007;25:461–475. 329. von Wolff M, Strowitzki T, Becker V, et al. Endometrial osteopontin, a ligand of β3-­integrin, is maximally expressed around the time of the “implantation” window. Fertil Steril. 2001;76:775–781. 330. Behzad F, Seif MW, Campbell S, et al. Expression of two isoforms of CD44 in human endometrium. Biol Reprod. 1994;51:739–747. 331. Lessey BA, Ilesanmi AO, Sun J, et al. Luminal and glandular endometrial epithelium express integrins differentially throughout the menstrual cycle: implications for implantation, contraception, and infertility. Am J Reprod Immunol. 1996;35:195–204. 332. Campbell S, Swann HR, Seif MW, et al. Cell. adhesion molecules on the oocyte and preimplantation human embryo. Hum Reprod. 1995;10:1571–1578. 333. Yelian FD, Edgeworth NA, Dong LJ, et al. Recombinant entactin promotes mouse primary trophoblast cell adhesion and migration through the Arg-­Gly-­Asp (RGD) recognition sequence. J Cell Biol. 1993;121:923–929. 334. Yelian FD, Yang Y, Hirata JD, et al. Molecular interactions between fibronectin and integrins during mouse blastocyst outgrowth. Mol Reprod Dev. 1995;41:435–448. 335. Raab G, Kover K, Paria BC, et al. Mouse preimplantation blastocysts adhere to cells expressing the transmembrane form of heparin-­ binding EGF-­like growth factor. Development. 1996;122:637–645. 336. Lai TH, Shih IM, Vlahos N, et al. Differential expression of L-­selectin ligand in the endometrium during the menstrual cycle. Fertil Steril. 2005;83(suppl 1):1297–1302. 337. Foulk RA, Zdravkovic T, Genbacev O, et al. Expression of L-­selectin ligand MECA-­79 as a predictive marker of human uterine receptivity. J Assist Reprod Genet. 2007;24:316–321. 338. Nikas G, Psychoyos A. Uterine pinopodes in peri-­ implantation human endometrium—Clinical relevance. Ann N Y Acad Sci. 1997;816:129–142. 339. Enders AC, Nelson DM. Pinocytotic activity of the uterus of the rat. Am J Anat. 1973;138:277–299. 340. Murphy CR. Understanding the apical surface markers of uterine receptivity. Pinopods or uterodomes? Hum Reprod. 2000;15:2451–2454. 341. Adams SM, Gayer N, Hosie MJ, et al. Human uterodomes (pinopods) do not display pinocytotic function. Hum Reprod. 2002;17:1980–1986. 342. Lessey BA, Gui Y, Apparao KB, et al. Regulated expression of heparin-­binding EGF-­like growth factor (HB-­EGF) in the human endometrium: a potential paracrine role during implantation. Mol Reprod Dev

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253.e8 PART I  The Fundamentals of Reproduction 343. Lessey BA. Two pathways of progesterone action in the human endometrium: implications for implantation and contraception. Steroids. 2003;68:809–815. 344. Kodaman PH, Taylor HS. Hormonal regulation of implantation. Obstet Gynecol Clin North Am. 2004;31:745–766 (ix). 345. Quinn CE, Detmar J, Casper RF. Pinopodes are present in Lif null and Hoxa10 null mice. Fertil Steril. 2007;88:1021–1028. 346. Usadi RS, Murray MJ, Bagnell RC, et al. Temporal and morphologic characteristics of pinopod expression across the secretory phase of the endometrial cycle in normally cycling women with proven fertility. Fertil Steril. 2003;79:970–974. 347. Acosta AA, Elberger L, Borghi M, et al. Endometrial dating and determination of the window of implantation in healthy fertile women. Fertil Steril. 2000;73:788–798. 348. Quinn C, Ryan E, Claessens EA, et al. The presence of pinopodes in the human endometrium does not delineate the implantation window. Fertil Steril. 2007;87:1015–1021. 349. Jones RL, Stoikos C, Findlay JK, et al. TGF-­beta superfamily expression and actions in the endometrium and placenta. Reproduction. 2006;132:217–232. 350. Norwitz ER, Schust DJ, Fisher SJ. Implantation and survival of early pregnancy. N Engl J Med. 2001;345:1400–1408. 351. Adams JC, Watt FM. Regulation of development and differentiation by the extracellular matrix. Development. 1993;117:1183–1198. 352. Turpeenniemi-­Hujanen T, Feinberg RF, Kauppila A, et al. Extracellular matrix interactions in early human embryos: implications for normal implantation events. Fertil Steril. 1995;64:132–138. 353. Santala P, Heino J. Regulation of integrin-­type cell adhesion receptors by cytokines. J Biol Chem. 1991;266:23505–23509. 354. Paiva P, Menkhorst E, Salamonsen L, et al. Leukemia inhibitory factor and interleukin-­11: critical regulators in the establishment of pregnancy. Cytokine Growth Factor Rev. 2009;20:319–328. 355. Cork BA, Tuckerman EM, Li TC, et al. Expression of interleukin (IL)-­11 receptor by the human endometrium in vivo and effects of IL-­11, IL-­6 and LIF on the production of MMP and cytokines by human endometrial cells in vitro. Mol Hum Reprod. 2002;8:841–848. 356. Cork BA, Li TC, Warren MA, et al. Interleukin-­11 (IL-­11) in human endometrium: expression throughout the menstrual cycle and the effects of cytokines on endometrial IL-­ 11 production in vitro. J Reprod Immunol. 2001;50:3–17. 357. Cullinan EB, Abbondanzo SJ, Anderson PS, et al. Leukemia inhibitory factor (LIF) and LIF receptor expression in human endometrium suggests a potential autocrine/paracrine function in regulating embryo implantation. Proc Natl Acad Sci U S A. 1996;93:3115–3120. 358. Lass A, Weiser W, Munafo A, et al. Leukemia inhibitory factor in human reproduction. Fertil Steril. 2001;76:1091–1096. 359. Stewart CL, Kaspar P, Brunet LJ, et al. Blastocyst implantation depends on maternal expression of leukaemia inhibitory factor. Nature. 1992;359:76–79. 360. Chen JR, Cheng JG, Shatzer T, et al. Leukemia inhibitory factor can substitute for nidatory estrogen and is essential to inducing a receptive uterus for implantation but is not essential for subsequent embryogenesis. Endocrinology. 2000;141:4365–4372. 361. Stewart CL. Leukaemia inhibitory factor and the regulation of pre-­ implantation development of the mammalian embryo. Mol Reprod Dev. 1994;39:233–238. 362. Dagoneau N, Scheffer D, Huber C, et al. Null leukemia inhibitory factor receptor (LIFR) mutations in Stuve-­Wiedemann/Schwartz-­ Jampel type 2 syndrome. Am J Hum Genet. 2004;74:298–305. 363. Nachtigall MJ, Kliman HJ, Feinberg RF, et al. The effect of leukemia inhibitory factor (LIF) on trophoblast differentiation: a potential role in human implantation. J Clin Endocrinol Metab. 1996;81:801–806. 364. Chaouat G, Menu E, Delage G, et al. Immuno-­endocrine interactions in early pregnancy. Hum Reprod. 1995;10(suppl 2):55–59. 365. Kelleher AM, Demayo FJ, Spencer TE. Uterine glands: developmental biology and functional roles in pregnancy. Endocr Rev. 2019;40(5):1424–1445. 366. Kelleher AM, Milano-­Foster J, Behura SK, Spencer TE. Uterine glands coordinate on-­time embryo implantation and impact endometrial decidualization for pregnancy success. Nat Commun. 2018;9(1). 367. Charnock-­Jones DS, Sharkey AM, Fenwick P, et al. Leukaemia inhibitory factor mRNA concentration peaks in human endometrium at the time of implantation and the blastocyst contains mRNA for the receptor at this time.

368. Arici A, Engin O, Attar E, et al. Modulation of leukemia inhibitory factor gene expression and protein biosynthesis in human endometrium. J Clin Endocrinol Metab. 1995;80:1908. 369. Chen DB, Hilsenrath R, Yang ZM, et al. Leukaemia inhibitory factor in human endometrium during the menstrual cycle: cellular origin and action on production of glandular epithelial cell prostaglandin in vitro. Hum Reprod. 1995;10:911. 370. Laird SM, Tuckerman EM, Dalton CF, et al. The production of leukaemia inhibitory factor by human endometrium: presence in uterine flushings and production by cells in culture. Hum Reprod. 1997;12:569. 371. Gonzalez RR, Rueda BR, Ramos MP, et al. Leptin-­induced increase in leukemia inhibitory factor and its receptor by human endometrium is partially mediated by interleukin 1 receptor signaling. Endocrinology. 2004;145:3850–3857. 372. Ernst M, Inglese M, Waring P, et al. Defective gp130-­mediated signal transducer and activator of transcription (STAT) signaling results in degenerative joint disease, gastrointestinal ulceration, and failure of uterine implantation. J Exp Med. 2001;194:189–203. 373. Song H, Lim H, Das SK, et al. Dysregulation of EGF family of growth factors and COX-­2 in the uterus during the preattachment and attachment reactions of the blastocyst with the luminal epithelium correlates with implantation failure in LIF-­deficient mice. Mol Endocrinol. 2000;14:1147–1161. 374. Fouladi-­Nashta AA, Jones CJ, Nijjar N, et al. Characterization of the uterine phenotype during the peri-­implantation period for LIF-­ null, MF1 strain mice. Dev Biol. 2005;281:1–21. 375. Jacobs AL, Carson DD. Uterine epithelial cell secretion of interleukin-­1 alpha induces prostaglandin E2 (PGE2) and PGF2 alpha secretion by uterine stromal cells in vitro. Endocrinology. 1993;132:300–308. 376. Jacobs AL, Hwang D, Julian J, et al. Regulated expression of prostaglandin endoperoxide synthase-­2 by uterine stroma. Endocrinology. 1994;135:1807–1815. 377. Lim H, Dey SK. PPAR delta functions as a prostacyclin receptor in blastocyst implantation. Trends Endocrinol Metab. 2000;11:137–142. 378. Lim H, Gupta RA, Ma WG, et al. Cyclo-­oxygenase-­2-­derived prostacyclin mediates embryo implantation in the mouse via PPARδ. Genes Dev. 1999;13:1561–1574. 379. Lim H, Paria BC, Das SK, et al. Multiple female reproductive failures in cyclooxygenase 2-­deficient mice. Cell. 1997;91:197–208. 380. Giess R, Tanasescu I, Steck T, et al. Leukaemia inhibitory factor gene mutations in infertile women. Mol Hum Reprod. 1999;5:581–586. 381. Robb L, Li RL, Hartley L, et al. Infertility in female mice lacking the receptor for interleukin 11 is due to a defective uterine response to implantation. Nat Med. 1998;4:303–308. 382. Dimitriadis E, Robb L, Salamonsen LA. Interleukin 11 advances progesterone-­induced decidualization of human endometrial stromal cells. Mol Hum Reprod. 2002;8:636–643. 383. Dimitriadis E, Stoikos C, Baca M, et al. Relaxin and prostaglandin E(2) regulate interleukin 11 during human endometrial stromal cell decidualization. J Clin Endocrinol Metab. 2005;90:3458–3465. 384. Dimitriadis E, Stoikos C, Tan YL, et al. Interleukin 11 signaling components signal transducer and activator of transcription 3 (STAT3) and suppressor of cytokine signaling 3 (SOCS3) regulate human endometrial stromal cell differentiation. Endocrinology. 2006;147:3809–3817. 385. Dimitriadis E, Sharkey AM, Tan YL, et al. Immunolocalisation of phosphorylated STAT3, interleukin 11 and leukaemia inhibitory factor in endometrium of women with unexplained infertility during the implantation window. Reprod Biol Endocrinol. 2007;5:44. 386. Smalley DM, Ley K. L-­selectin: mechanisms and physiological significance of ectodomain cleavage. J Cell Mol Med. 2005;9:255–266. 387. Thathiah A, Blobel CP, Carson DD. Tumor necrosis factor-­alpha converting enzyme/ADAM 17 mediates MUC1 shedding. J Biol Chem. 2003;278:3386–3394. 388. Thathiah A, Carson DD. MT1-­MMP mediates MUC1 shedding independent of TACE/ADAM17. Biochem J. 2004;382:363–373. 389. Franco HL, Lee KY, Broaddus RR, et al. Ablation of Indian hedgehog in the murine uterus results in decreased cell cycle progression, aberrant epidermal growth factor signaling, and increased estrogen signaling. Biol Reprod. 2010;82:783–790. 390. Roskoski Jr R. The ErbB/HER receptor protein-­tyrosine kinases and cancer. Biochem Biophys Res Commun. 2004;319:1–11. 391. Dey SK, Lim H, Das SK, et al. Molecular cues to implanta-

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract 392. Yoo HJ, Barlow DH, Mardon HJ. Temporal and spatial regulation of expression of heparin-­binding epidermal growth factor-­like growth factor in the human endometrium: a possible role in blastocyst implantation. Dev Genet. 1997;21:102–108. 393. Leach RE, Kilburn B, Wang J, et al. Heparin-­binding EGF-­like growth factor regulates human extravillous cytotrophoblast development during conversion to the invasive phenotype. Dev Biol. 2004;266:223–237. 394. Das SK, Wang XN, Paria BC, et al. Heparin-­binding EGF-­like growth factor gene is induced in the mouse uterus temporally by the blastocyst solely at the site of its apposition: a possible ligand for interaction with blastocyst EGF-­ receptor in implantation. Development. 1994;120:1071–1083. 395. Sporn MB, Roberts AB, Wakefield LM, et al. Some recent advances in the chemistry and biology of transforming growth factor-­beta. J Cell Biol. 1987;105:1039–1045. 396. Sporn MB, Roberts AB, Wakefield LM, et al. Transforming growth factor-­β: biological function and chemical structure. Science. 1986;233:532–534. 397. Godkin JD, Dore JJ. Transforming growth factor beta and the endometrium. Rev Reprod. 1998;3:1–6. 398. Kulkarni AB, Thyagarajan T, Letterio JJ. Function of cytokines within the TGF-­ beta superfamily as determined from transgenic and gene knockout studies in mice. Curr Mol Med. 2002;2:303–327. 399. Chegini N, Zhao Y, Williams RS, et al. Human uterine tissue throughout the menstrual cycle expresses transforming growth factor-­ beta 1 (TGF beta⋅1), TGF beta⋅2, TGF beta⋅3, and TGF⋅beta type II receptor messenger ribonucleic acid and protein and contains [125 I]TGF⋅1-­binding sites. Endocrinology. 1994;135:439–449. 400. Gold LI, Saxena B, Mittal KR, et al. Increased expression of transforming growth factor beta isoforms and basic fibroblast growth factor in complex hyperplasia and adenocarcinoma of the endometrium: evidence for paracrine and autocrine action. Cancer Res. 1994;54:2347–2358. 401. Gaide Chevronnay HP, Cornet PB, Delvaux D, et al. Opposite regulation of transforming growth factors-­beta 2 and -­beta 3 expression in the human endometrium. Endocrinology. 2007;149. 402. Gaide Chevronnay HP, Cornet PB, Delvaux D, et al. Opposite regulation of transforming growth factors-­beta2 and -­beta3 expression in the human endometrium. Endocrinology. 2008;149:1015–1025. 403. Bruner KL, Rodgers WH, Gold LI, et al. Transforming growth factor beta mediates the progesterone suppression of an epithelial metalloproteinase by adjacent stroma in the human endometrium. Proc Natl Acad Sci U S A. 1995;92:7362–7366. 404. Tang M, Taylor HS, Tabibzadeh S. In vivo gene transfer of lefty leads to implantation failure in mice. Hum Reprod. 2005;20:1772–1778. 405. Tierney EP, Giudice LC. Role of activin A as a mediator of in vitro endometrial stromal cell decidualization via the cyclic adenosine monophosphate pathway. Fertil Steril. 2004;81(suppl 1):899–903. 406. Kauma S, Matt D, Strom S, et al. Interleukin-­1 beta, human leukocyte antigen HLA-­DR alpha, and transforming growth factor-­beta expression in endometrium, placenta, and placental membranes. Am J Obstet Gynecol. 1990;163:1430–1437. 407. Feinberg RF, Kliman HJ, Wang CL. Transforming growth factor-­β stimulates trophoblast oncofetal fibronectin synthesis in vitro: implications for trophoblast implantation in vivo. J Clin Endocrinol Metab. 1994;78:1241–1248. 408. Ignotz RA, Massague J. Transforming growth factor-­beta stimulates the expression of fibronectin and collagen and their incorporation into the extracellular matrix. J Biol Chem. 1986;261:4337–4345. 409. Ng YH, Zhu H, Pallen CJ, et al. Differential effects of interleukin-­ 1beta and transforming growth factor-­beta1 on the expression of the inflammation-­associated protein, ADAMTS-­1, in human decidual stromal cells in vitro. Hum Reprod. 2006;21:1990–1999. 410. Irwin JC, de las Fuentes L, Giudice LC. Growth factors and decidualization in vitro. Ann N Y Acad Sci. 1994;734:7–18. 411. Fujimoto J, Hori M, Ichigo S, et al. Expression of basic fibroblast growth factor and its mRNA in uterine endometrium during the menstrual cycle. Gynecol Endocrinol. 1996;10:193–197. 412. Tsai SJ, Wu MH, Chen HM, et al. Fibroblast growth factor-­ 9 is an endometrial stromal growth factor. Endocrinology. 2002;143:2715–2721.

253.e9

413. Clemmons DR, Underwood LE. Clinical review 59: uses of human insulin-­like growth factor-­I in clinical conditions. J Clin Endocrinol Metab. 1994;79:4–6. 414. Giudice LC, Lamson G, Rosenfeld RG, et al. Insulin-­like growth factor-­II (IGF-­II) and IGF binding proteins in human endometrium. Ann N Y Acad Sci. 1991;626:295–307. 415. Zhou J, Dsupin BA, Giudice LC, et al. Insulin-­like growth factor system gene expression in human endometrium during the menstrual cycle. J Clin Endocrinol Metab. 1994;79:1723–1734. 416. Laatikainen TJ, Tomas EI, Voutilainen RJ. The expression of insulin-­like growth factor and its binding protein mRNA in the endometrium of postmenopausal patients with breast cancer receiving tamoxifen. Cancer. 1995;76:1406–1410. 417. Hild-­Petito S, Verhage HG, Fazleabas AT. Characterization, localization, and regulation of receptors for insulin-­like growth factor I in the baboon uterus during the cycle and pregnancy. Biol Reprod. 1994;50:791–801. 418. Tazuke SI, Giudice LC. Growth factors and cytokines in endometrium, embryonic development, and maternal-­embryonic interactions. Semin Reprod Endocrinol. 1996;14:231–245. 419. Giudice LC. Multifaceted roles for IGFBP-­ 1 in human endometrium during implantation and pregnancy. Ann N Y Acad Sci. 1997;828:146–156. 420. Hunt JS, Chen HL, Hu XL, et al. Tumor necrosis factor-­alpha messenger ribonucleic acid and protein in human endometrium. Biol Reprod. 1992;47:141–147. 421. Hampson J, McLaughlin PJ, Johnson PM. Low-­affinity receptors for tumour necrosis factor-­ alpha, interferon-­ gamma and granulocyte-­ macrophage colony-­ stimulating factor are expressed on human placental syncytiotrophoblast. Immunology. 1993;79:485–490. 422. Todt JC, Yang Y, Lei J. Effects of tumor necrosis factor-­alpha on human trophoblast cell adhesion and motility. Am J Reprod Immunol. 1996;36:65–71. 423. Grosskinsky CM, Yowell CW, Sun J, et al. Modulation of integrin expression in endometrial stromal cells in vitro. J Clin Endocrinol Metab. 1996;81:2047–2054. 424. Burgess AW, Wilson EM, Metcalf D. Stimulation by human placental conditioned medium of hemopoietic colony formation by human marrow cells. Blood. 1977;49:573–583. 425. Pollard JW, Hunt JS, Wiktor-­Jedrzejczak W, et al. A pregnancy defect in the osteopetrotic (op/op) mouse demonstrates the requirement for CSF-­1 in female fertility. Dev Biol. 1991;148:273–283. 426. Robertson SA, Roberts CT, Farr KL, et al. Fertility impairment in granulocyte-­macrophage colony-­stimulating factor-­deficient mice. Biol Reprod. 1999;60:251–261. 427. Daiter E, Pampfer S, Yeung YG, et al. Expression of colony-­ stimulating factor-­ 1 in the human uterus and placenta. J Clin Endocrinol Metab. 1992;74:850–858. 428. Omigbodun A, Coukos G, Ziolkiewicz P, et al. Macrophage-­ colony stimulating factor (M-­ CSF) regulates the expression of fibronectin and its alpha5 integrin receptor in human trophoblasts. Endocrinology. 1998;139:2190–2193. 429. Hansen PJ, Dobbs KB, Denicol AC, et al. Sex and the preimplantation embryo: implications of sexual dimorphism in the preimplantation period for maternal programming of embryonic development. Cell Tissue Res. 2016;363:237–247. 430. Aleyasin A, Abediasl Z, Nazari A, et al. Granulocyte colony-­ stimulating factor in repeated IVF failure, a randomized trial. Reproduction. 2016;151:637–642. 431. Scarpellini F, Sbracia M. Use of granulocyte colony-­stimulating factor for the treatment of unexplained recurrent miscarriage: a randomised controlled trial. Hum Reprod. 2009;24:2703–2708. 432. Stewart EA. Gonadotropins and the uterus: is there a gonad-­ independent pathway? J Soc Gynecol Investig. 2001;8:319–326. 433. Lobo SC, Srisuparp S, Peng X, et al. Uterine receptivity in the baboon: modulation by chorionic gonadotropin. Semin Reprod Med. 2001;19:69–74. 434. Fanchin R, Peltier E, Frydman R, et al. Human chorionic gonadotropin: does it affect human endometrial morphology in vivo? Semin Reprod Med. 2001;19:31–35. 435. Lovely LP, Fazleabas AT, Fritz MA, et al. Prevention of endometrial apoptosis: randomized prospective comparison of human chorionic gonadotropin versus progesterone treatment in the luteal . 2005;90:2351–2356.

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253.e10 PART I  The Fundamentals of Reproduction 436. Fox C, Azores-­Gococo D, Swart L, et al. Luteal phase hCG support for unexplained recurrent pregnancy loss—A low hanging fruit? Reprod Biomed Online. 2017;34(3):319–324. 437. Sherwin JR, Hastings JM, Jackson KS, et al. The endometrial response to chorionic gonadotropin is blunted in a baboon model of endometriosis. Endocrinology. 2010;151:4982–4993. 438. Strakova Z, Mavrogianis P, Meng X, et al. In vivo infusion of interleukin-­ 1beta and chorionic gonadotropin induces endometrial changes that mimic early pregnancy events in the baboon. Endocrinology. 2005;146:4097–4104. 439. Strug MR, Su R, Young JE, et al. Intrauterine human chorionic gonadotropin infusion in oocyte donors promotes endometrial synchrony and induction of early decidual markers for stromal survival: a randomized clinical trial. Hum Reprod. 2016;31:1552–1561. 440. Fluhr H, Carli S, Deperschmidt M, et al. Differential effects of human chorionic gonadotropin and decidualization on insulin-­like growth factors-­I and -­II in human endometrial stromal cells. Fertil Steril. 2008;90(suppl 4):1384–1389. 441. Sugihara K, Kabir-­Salmani M, Byrne J, et al. Induction of trophinin in human endometrial surface epithelia by CGbeta and IL-­1beta. FEBS Lett. 2007;582:197–202. 442. Sherwin JR, Sharkey AM, Cameo P, et al. Identification of novel genes regulated by chorionic gonadotropin in baboon endometrium during the window of implantation. Endocrinology. 2007;148:618–626. 443. Marions L, Danielsson KG. Expression of cyclo-­ oxygenase in human endometrium during the implantation period. Mol Hum Reprod. 1999;5:961–965. 444. Li DK, Liu L, Odouli R. Exposure to non-­ steroidal anti-­ inflammatory drugs during pregnancy and risk of miscarriage: population based cohort study. BMJ. 2003;327:368. 445. Lim H, Dey SK. A novel pathway of prostacyclin signaling-­hanging out with nuclear receptors. Endocrinology. 2002;143:3207–3210. 446. Ding NZ, Ma XH, Diao HL, et al. Differential expression of peroxisome proliferator-­activated receptor delta at implantation sites and in decidual cells of rat uterus. Reproduction. 2003;125:817–825. 447. Matsumoto H, Ma W, Smalley W, et al. Diversification of cyclooxygenase-­2-­derived prostaglandins in ovulation and implantation. Biol Reprod. 2001;64:1557–1565. 448. Paria BC, Wang H, Dey SK. Endocannabinoid signaling in synchronizing embryo development and uterine receptivity for implantation. Chem Phys Lipids. 2002;121:201–210. 449. Habayeb OM, Bell SC, Konje JC. Endogenous cannabinoids: metabolism and their role in reproduction. Life Sci. 2002;70:1963–1977. 450. Schuel H, Burkman LJ, Lippes J, et al. N-­Acylethanolamines in human reproductive fluids. Chem Phys Lipids. 2002;121:211–227. 451. Maccarrone M, Bisogno T, Valensise H, et al. Low fatty acid amide hydrolase and high anandamide levels are associated with failure to achieve an ongoing pregnancy after IVF and embryo transfer. Mol Hum Reprod. 2002;8:188–195. 452. Barash A, Dekel N, Fieldust S, Segal I, Schechtman E, Granot I. Local injury to the endometrium doubles the incidence of successful pregnancies in patients undergoing in vitro fertilization. Fertil Steril. 2003;79(6):1317–1322. 453. Metwally M, Walters S, Chatters R. The current role of induced endometrial trauma (endometrial scratch) in women undergoing infertility treatment. Semin Reprod Med. 2021;39(5–06):e1–e4. 454. Robb CT, Regan KH, Dorward DA, et al. Key mechanisms governing resolution of lung inflammation. Semin Immunopathol. 2016;38:425–448. 455. Canny GO, Lessey BA. The role of lipoxin A4 in endometrial biology and endometriosis. Mucosal Immunol. 2013;6(3):439–450. 456. Hyde KJ, Schust DJ. Immunologic challenges of human reproduction: an evolving story. Fertil Steril. 2016;106:499–510. 457. Whitelaw PF, Croy BA. Granulated lymphocytes of pregnancy. Placenta. 1996;17:533–543. 458. Lea RG, Sandra O. Immunoendocrine aspects of endometrial function and implantation. Reproduction. 2007;134:389–404. 459. Givan AL, White HD, Stern JE, et al. Flow cytometric analysis of leukocytes in the human female reproductive tract: comparison of fallopian tube, uterus, cervix, and vagina. Am J Reprod Immunol. 1997;38:350–359. 460. Hannan NJ, Salamonsen LA. Role of chemokines in the endometrium and in embryo implantation. Curr Opin Obstet Gynecol. 2007;19:266–272.

461. Ma Z, Lu FX, Torten M, et al. The number and distribution of immune cells in the cervicovaginal mucosa remain constant throughout the menstrual cycle of rhesus macaques. Clin Immunol. 2001;100:240–249. 462. Andrews AL, Nasir T, Bucchieri F, et al. IL-­13 receptor alpha 2: a regulator of IL-­13 and IL-­4 signal transduction in primary human fibroblasts. J Allergy Clin Immunol. 2006;118:858–865. 463. Szekeres-­Bartho J, Halasz M, Palkovics T. Progesterone in pregnancy; receptor-­ligand interaction and signaling pathways. J Reprod Immunol. 2009;83:60–64. 464. Bulmer JN, Longfellow M, Ritson A. Leukocytes and resident blood cells in endometrium. Ann N Y Acad Sci. 1991;622:57. 465. Bulmer JN, Morrison I, Longfellow M, et al. Granulated lymphocytes in human endometrium; histological and immunohistochemical studies. Hum Reprod. 1991;6:791–798. 466. Pace D, Longfellow M, Bulmer JN. Characterization of intraepithelial lymphocytes in human endometrium. J Reprod Fertil. 1991;91:165–174. 467. Jones RL, Kelly RW, Critchley HOD. Chemokine and cyclooxygenase-­2 expression in human endometrium coincides with leukocyte accumulation. Hum Reprod. 1997;12:1300. 468. Kelly RW, King AE, Critchley HO. Cytokine control in human endometrium. Reproduction. 2001;121:3–19. 469. Dimitriadis E, White CA, Jones RL, et al. Cytokines, chemokines and growth factors in endometrium related to implantation. Hum Reprod Update. 2005;11:613–630. 470. Kitaya K, Yamaguchi T, Honjo H. Central role of interleukin-­15 in postovulatory recruitment of peripheral blood CD16(-­) natural killer cells into human endometrium. J Clin Endocrinol Metab. 2005;90:2932–2940. 471. Shifren JL, Tseng JF, Zaloudek CJ, et al. Ovarian steroid regulation of vascular endothelial growth factor in the human endometrium: implications for angiogenesis during the menstrual cycle and in the pathogenesis of endometriosis. J Clin Endocrinol Metab. 1996;81:3112. 472. Kennedy TG, Lukash LA. Induction of decidualization in rats by the intrauterine infusion of prostaglandins. Biol Reprod. 1982;27:253–260. 473. St-­Louis I, Singh M, Brasseur K, et al. Expression of COX-­1 and COX-­2 in the endometrium of cyclic, pregnant and in a model of pseudopregnant rats and their regulation by sex steroids. Reprod Biol Endocrinol. 2010;8:103. 474. Poyser NL. The control of prostaglandin production by the endometrium in relation to luteolysis and menstruation. Prostaglandins Leukot Essent Fatty Acids. 1995;53:147. 475. Frank GR, Brar AK, Cedars MI, et al. Prostaglandin E2 enhances human endometrial stromal cell differentiation. Endocrinology. 1994;134:258. 476. Achache H, Tsafrir A, Prus D, et al. Defective endometrial prostaglandin synthesis identified in patients with repeated implantation failure undergoing in vitro fertilization. Fertil Steril. 2010;94(4):1271–1278. 477. Wang Y, Zhao AM, Lin QD. Role of cyclooxygenase-­2 signaling pathway dysfunction in unexplained recurrent spontaneous abortion. Chin Med J. 2010;123:1543–1547. 478. Laird SM, Tuckerman EM, Cork BA, et al. A review of immune cells and molecules in women with recurrent miscarriage. Hum Reprod Update. 2003;9:163–174. 479. Jasper MJ, Tremellen KP, Robertson SA. Primary unexplained infertility is associated with reduced expression of the T-­regulatory cell transcription factor Foxp3 in endometrial tissue. Mol Hum Reprod. 2006;12:301–308. 480. Quenby S, Nik H, Innes B, et al. Uterine natural killer cells and angiogenesis in recurrent reproductive failure. Hum Reprod. 2009;24:45–54. 481. Johnson PM, Christmas SE, Vince GS. Immunological aspects of implantation and implantation failure. Hum Reprod. 1999;14(suppl 2):26. 482. Baulieu EE. Contragestion and other clinical applications of RU-­ 486, an antiprogesterone at the receptor. Science. 1989;245:1351–1357. 483. van Mourik MS, Macklon NS, Heijnen CJ. Embryonic implantation: cytokines, adhesion molecules, and immune cells in establishing an implantation environment. J Leukoc Biol. 2009;85:

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract253.e11 484. Robertson SA, Guerin LR, Moldenhauer LM, et al. Activating T regulatory cells for tolerance in early pregnancy—The contribution of seminal fluid. J Reprod Immunol. 2009;83:109–116. 485. Pavone ME, Reierstad S, Sun H, et al. Altered retinoid uptake and action contributes to cell survival in endometriosis. J Clin Endocrinol Metab. 2010;95:E300–E309. 486. Bulun SE, Cheng YH, Pavone ME, et al. 17Beta-­hydroxysteroid dehydrogenase-­2 deficiency and progesterone resistance in endometriosis. Semin Reprod Med. 2010;28:44–50. 487. Mucida D, Pino-­Lagos K, Kim G, et al. Retinoic acid can directly promote TGF-­β-­mediated Foxp3(+) Treg cell conversion of naive T cells. Immunity. 2009;30:471–472. , author reply 472-­473. 488. Guerin LR, Prins JR, Robertson SA. Regulatory T-­ cells and immune tolerance in pregnancy: a new target for infertility treatment? Hum Reprod Update. 2009;15:517–535. 489. Jasper MJ, Care AS, Sullivan B, et al. Macrophage-­derived LIF and IL1B regulate alpha(1,2)fucosyltransferase 2 (Fut2) expression in mouse uterine epithelial cells during early pregnancy. Biol Reprod. 2011;84:179–188. 490. Nakamura H, Jasper MJ, Hull ML, et al. Macrophages regulate expression of α1,2-­fucosyltransferase genes in human endometrial epithelial cells. Mol Hum Reprod. 2012;18:204–215. 491. Na YJ, Jin JO, Lee MS, et al. Peritoneal fluid from endometriosis patients switches differentiation of monocytes from dendritic cells to macrophages. J Reprod Immunol. 2008;77:63–74. 492. Kamei M, Carman CV. New observations on the trafficking and diapedesis of monocytes. Curr Opin Hematol. 2010;17:43–52. 493. Du H, Taylor HS. Contribution of bone marrow-­derived stem cells to endometrium and endometriosis. Stem Cell. 2007;25:2082–2086. 494. Bulmer JN, Lash GE. Human natural killer cells: a reappraisal. Mol Immunol. 2005;42:511–521. 495. Collins MK, Tay CS, Erlebacher A. Dendritic cell entrapment within the pregnant uterus inhibits immune surveillance of the maternal/fetal interface in mice. J Clin Invest. 2009;119:2062–2073. 496. Hanna J, Mandelboim O. When killers become helpers. Trends Immunol. 2007;28:201–206. 497. Kalkunte S, Chichester CO, Gotsch F, et al. Evolution of non-­ cytotoxic uterine natural killer cells. Am J Reprod Immunol. 2008;59:425–432. 498. Kämmerer U, Schoppet M, McLellan AD, et al. Human decidua contains potent immunostimulatory CD83+ dendritic cells. Am J Pathol. 2000;157:159. 499. Lea RG, Clark DA. Macrophages and migratory cells in endometrium relevant to implantation. Baillieres Clin Obstet Gynaecol. 1991;5:25–59. 500. Oh MJ, Croy BA. A map of relationships between uterine natural killer cells and progesterone receptor expressing cells during mouse pregnancy. Placenta. 2008;29:317–323. 501. Oliver C, Cowdrey N, Abadia-­Molina AC, et al. Antigen phenotype of cultured decidual stromal cells of human term decidua. J Reprod Immunol. 1999;45:19–30. 502. Sanguansermsri D, Pongcharoen S. Pregnancy immunology: decidual immune cells. Asian Pac J Allergy Immunol. 2008;26:171–181. 503. Yoshinaga K. Review of factors essential for blastocyst implantation for their modulating effects on the maternal immune system. Semin Cell Dev Biol. 2008;19:161–169. 504. Vivier E, Raulet DH, Moretta A, et al. Innate or adaptive immunity? The example of natural killer cells. Science. 2011;331:44–49. 505. Aluvihare VR, Kallikourdis M, Betz AG. Regulatory T cells mediate maternal tolerance to the fetus. Nat Immunol. 2004;5:266–271. 506. Saito S, Sasaki Y, Sakai M. CD4(+)CD25high regulatory T cells in human pregnancy. J Reprod Immunol. 2005;65:111–120. 507. Zenclussen AC. CD4(+)CD25+ T regulatory cells in murine pregnancy. J Reprod Immunol. 2005;65:101–110. 508. Aluvihare VR, Betz AG. The role of regulatory T cells in alloantigen tolerance. Immunol Rev. 2006;212:330–343. 509. Sakaguchi S. Regulatory T cells: key controllers of immunologic self-­tolerance. Cell. 2000;101:455–458. 510. Robertson SA, Guerin LR, Bromfield JJ, et al. Seminal fluid drives expansion of the CD4+CD25+ T regulatory cell pool and induces tolerance to paternal alloantigens in mice. Biol Reprod. 2009;80:1036–1045. 511. Mai J, Wang H, Yang XF. Th 17 cells interplay with Foxp3+ Tregs in regulation of inflammation and autoimmunity. 2010;15:986–1006.

512. Elyaman W, Bradshaw EM, Uyttenhove C, et al. IL-­9 induces differentiation of TH17 cells and enhances function of FoxP3+ natural regulatory T cells. Proc Natl Acad Sci U S A. 2009;106:12885–12890. 513. Nowak EC, Weaver CT, Turner H, et al. IL-­9 as a mediator of Th17-­driven inflammatory disease. J Exp Med. 2009;206:1653–1660. 514. Jabbour HN, Critchley HO. Potential roles of decidual prolactin in early pregnancy. Reproduction. 2001;121:197–205. 515. Dosiou C, Giudice LC. Natural killer cells in pregnancy and recurrent pregnancy loss: endocrine and immunologic perspectives. Endocr Rev. 2005;26:44–62. 516. Doisne JM, Balmas E, Boulenouar S, et al. Composition, development, and function of uterine innate lymphoid cells. J Immunol. 2015;195:3937–3945. 517. Vacca P, Montaldo E, Croxatto D, et al. Identification of diverse innate lymphoid cells in human decidua. Mucosal Immunol. 2015;8:254–264. 518. Buonocore S, Ahern PP, Uhlig HH, et al. Innate lymphoid cells drive interleukin-­23-­dependent innate intestinal pathology. Nature. 2010;464:1371–1375. 519. Brackin MN, Cruse JM, Lewis RE, et al. Quantitative analysis of adhesion molecules on cellular constituents of the human uterine microenvironment under the influence of estrogen and progesterone. Exp Mol Pathol. 2002;72:91–114. 520. Piccinni MP, Beloni L, Livi C, et al. Defective production of both leukemia inhibitory factor and type 2 T-­ helper cytokines by decidual T cells in unexplained recurrent abortions. Nat Med. 1998;4:1020–1024. 521. Fraccaroli L, Grasso E, Zeitler E, et al. Modulation of maternal LIF producers T cells by trophoblast and paternal antigens. Am. J Reprod Immunol. 2011;65:133–145. 522. Illera MJ, Juan L, Stewart CL, et al. Effect of peritoneal fluid from women with endometriosis on implantation in the mouse model. Fertil Steril. 2000;74:41–48. 523. Aghajanova L, Altmae S, Bjuresten K, et al. Disturbances in the LIF pathway in the endometrium among women with unexplained infertility. Fertil Steril. 2009;91:2602–2610. 524. Li L, Xu BF, Chen QJ, et al. Effects of hydrosalpinx on pinopodes, leukaemia inhibitory factor, integrin beta3 and MUC1 expression in the peri-­implantation endometrium. Eur J Obstet Gynecol Reprod Biol. 2010;151:171–175. 525. Zhao M, Chang C, Liu Z, et al. Treatment with low-­dose aspirin increased the level LIF and integrin beta3 expression in mice during the implantation window. Placenta. 2010;31:1101–1105. 526. Franasiak J, Holoch K, Yuan L, et al. Leukemia inhibitory factor and anb3 integrin expression in normal and abnormal endometrium. Fertil Steril. 2012 (in preparation). 527. Miller PB, Parnell BA, Bushnell G, et al. Endometrial receptivity defects during IVF cycles with and without letrozole. Hum Reprod. 2012;27:881–888. 528. Ledee-­Bataille N, Lapree-­Delage G, Taupin JL, et al. Concentration of leukaemia inhibitory factor (LIF) in uterine flushing fluid is highly predictive of embryo implantation. Hum Reprod. 2002;17:213–218. 529. Serafini PC, Silva ID, Smith GD, et al. Endometrial claudin-­4 and leukemia inhibitory factor are associated with assisted reproduction outcome. Reprod Biol Endocrinol. 2009;7:30. 530. Lessey BA, Castelbaum AJ, Sawin SW, et al. Aberrant integrin expression in the endometrium of women with endometriosis. J Clin Endocrinol Metab. 1994;79:643–649. 531. Meyer WR, Castelbaum AJ, Somkuti S, et al. Hydrosalpinges adversely affect markers of endometrial receptivity. Hum Reprod. 1997;12:1393–1398. 532. Weber MA, Schnyder-­Candrian S, Schnyder B, et al. Endogenous leukemia inhibitory factor attenuates endotoxin response. Lab Invest. 2005;85:276–284. 533. Waring PM, Waring LJ, Billington T, et al. Leukemia inhibitory factor protects against experimental lethal Escherichia coli septic shock in mice. Proc Natl Acad Sci U S A. 1995;92:1337–1341. 534. Waring P, Wycherley K, Cary D, et al. Leukemia inhibitory factor levels are elevated in septic shock and various inflammatory body fluids. J Clin Invest. 1992;90:2031–2037. 535. Young SL, Lessey BA, Murray MJ, et al. Regulation and Function of Human Endometrial Decay Accelerating Factor (DAF). 47th Annual Meeting of the Society of Gynecologic Investigation; 2000. 536. Fedarko NS, Jain A, Karadag A, et al. Three small integrin binding linked glycoproteins (SIBLINGs) bind and activate specific matrix metalloproteinases. FASEB J. 2004;18:734–736.

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253.e12 PART I  The Fundamentals of Reproduction 537. Piccinni MP, Giudizi MG, Biagiotti R, et al. Progesterone favors the development of human T helper cells producing Th2-­type cytokines and promotes both IL-­ 4 production and membrane CD30 expression in established Th1 cell clones. J Immunol. 1995;155:128–133. 538. Hasty LA, Lambris JD, Lessey BA, et al. Hormonal regulation of complement components and receptors throughout the menstrual cycle. Am J Obstet Gynecol. 1994;170:168–175. 539. Isaacson KB, Galman M, Coutifaris C, et al. Endometrial synthesis and secretion of complement component-­3 by patients with and without endometriosis. Fertil Steril. 1990;53:836–841. 540. Jensen TS, Bjorge L, Wollen AL, et al. Identification of the complement regulatory proteins CD46, CD55, and CD59 in human fallopian tube, endometrium, and cervical mucosa and secretion. Am J Reprod Immunol. 1995;34:1–9. 541. Young SL, Lessey BA, Fritz MA, et al. In vivo and in vitro evidence suggest that HB-­ EGF regulates endometrial expression of human decay-­ accelerating factor. J Clin Endocrinol Metab. 2002;87:1368–1375. 542. Xu C, Mao D, Holers VM, et al. A critical role for murine complement regulator crry in fetomaternal tolerance. Science. 2000;287:498–501. 543. Holers VM, Girardi G, Mo L, et al. Complement C3 activation is required for antiphospholipid antibody-­induced fetal loss. J Exp Med. 2002;195:211–220. 544. Salmon JE, Girardi G, Holers VM. Complement activation as a mediator of antiphospholipid antibody induced pregnancy loss and thrombosis. Ann Rheum Dis. 2003;61(suppl 2):ii46–ii50. 545. Thurman JM, Kraus DM, Girardi G, et al. A novel inhibitor of the alternative complement pathway prevents antiphosphoslipid antibody-­ induced pregnancy loss in mice. Mol Immunol. 2005; 42:87–97. 546. King AE, Fleming DC, Critchley HO, et al. Regulation of natural antibiotic expression by inflammatory mediators and mimics of infection in human endometrial epithelial cells. Mol Hum Reprod. 2002;8:341–349. 547. Quayle AJ, Porter EM, Nussbaum AA, et al. Gene expression, immunolocalization, and secretion of human defensin-­5 in human female reproductive tract. Am J Pathol. 1998;152:1247–1258. 548. Maranduba CM da C, De Castro SBR, de Souza GT, et al. Intestinal microbiota as modulators of the immune system and neuroimmune system: impact on the host health and homeostasis. J Immunol Res. 2015;2015:931574. 549. Flint HJ, Scott KP, Louis P, Duncan SH. The role of the gut microbiota in nutrition and health. Nat Rev Gastroenterol Hepatol. 2012;9(10):577–589. 550. Chen C, Song X, Wei W, et al. The microbiota continuum along the female reproductive tract and its relation to uterine-­related diseases. Nat Commun. 2017;8(1):875. 551. Moreno I, Codoñer FM, Vilella F, et al. Evidence that the endometrial microbiota has an effect on implantation success or failure. Am J Obstet Gynecol. 2016;215(6):684–703. 552. Franasiak JM, Werner MD, Juneau CR, et al. Endometrial microbiome at the time of embryo transfer: next-­generation sequencing of the 16S ribosomal subunit. J Assist Reprod Genet. 2016;33(1):129–136. 553. Riganelli L, Iebba V, Piccioni M, et al. Structural variations of vaginal and endometrial Microbiota: Hints on female infertility. Front Cell Infect Microbiol. 2020;10:350. 554. Noyes RW, Hertig AI, Rock J. Dating the endometrial biopsy. Fertil Steril. 1950;1:3–25. 555. Coutifaris C, Myers ER, Guzick DS, et al. Histological dating of timed endometrial biopsy tissue is not related to fertility status. Fertil Steril. 2004;82:1264–1272. 556. Murray MJ, Meyer WR, Zaino RJ, et al. A critical analysis of the accuracy, reproducibility, and clinical utility of histologic endometrial dating in fertile women. Fertil Steril. 2004;81:1333–1343. 557. Fritz MA, Lessey BA. Defective luteal function. In: Fraser IS, Jansen RPS, Lobo RA, et al., eds. Estrogens and progestogens in clinical practice. vol 1. London: Churchill Livingstone; 1998:437–453. 558.  Practice Committee of American Society for Reproductive M. Current clinical irrelevance of luteal phase deficiency: a committee opinion. Fertil Steril. 2012;98:1112–1117. 559. Prapas Y, Prapas N, Jones EE, et al. The window for embryo transfer in oocyte donation cycles depends on the duration of progesterone therapy. Hum Reprod

560. Wilcox AJ, Weinberg CR, Baird DD. Timing of sexual intercourse in relation to ovulation. Effects on the probability of conception, survival of the pregnancy, and sex of the baby. N Engl J Med. 1995;333(23):1517–1521. 561. Young SL, Savaris RF, Lessey BA, et al. Effect of randomized serum progesterone concentration on secretory endometrial histologic development and gene expression. Hum Reprod. 2017;32(9):1903–1914. 562. Barash A, Dekel N, Fieldust S, et al. Local injury to the endometrium doubles the incidence of successful pregnancies in patients undergoing in vitro fertilization. Fertil Steril. 2003;79:1317–1322. 563. Zhou L, Li R, Wang R, et al. Local injury to the endometrium in controlled ovarian hyperstimulation cycles improves implantation rates. Fertil Steril. 2008;89:1166–1176. 564. Karimzadeh MA, Ayazi Rozbahani M, Tabibnejad N. Endometrial local injury improves the pregnancy rate among recurrent implantation failure patients undergoing in vitro fertilisation/intra cytoplasmic sperm injection: a randomised clinical trial. Aust N Z J Obstet Gynaecol. 2009;49:677–680. 565. El-­Toukhy T, Sunkara S, Khalaf Y. Local endometrial injury and IVF outcome: a systematic review and meta-­analysis. Reprod Biomed Online. 2012;25:345–354. 566. Panagiotopoulou N, Karavolos S, Choudhary M. Endometrial injury prior to assisted reproductive techniques for recurrent implantation failure: a systematic literature review. Eur J Obstet Gynecol Reprod Biol. 2015;193:27–33. 567. Gnainsky Y, Granot I, Aldo PB, et al. Local injury of the endometrium induces an inflammatory response that promotes successful implantation. Fertil Steril. 2010;94:2030–2036. 568. Dekel N, Gnainsky Y, Granot I, et al. The role of inflammation for a successful implantation. Am. J Reprod Immunol. 2014;72:141–147. 569. Gnainsky Y, Granot I, Aldo P, et al. Biopsy-­induced inflammatory conditions improve endometrial receptivity: the mechanism of action. Reproduction. 2015;149:75–85. 570. Lessey BA, Gardner DK, Weissman A, et al. The use of biomarkers for the assessment of uterine receptivity. In: Gardner DK, et al., ed. Textbook of Assisted Reproductive Technology. Martin Dunitz Ltd; 2001:353. 571. Cavagna M, Mantese JC. Biomarkers of endometrial receptivity—A review. Placenta. 2003;24(suppl B):S39–S47. 572. Sha AG, Liu JL, Jiang XM, et al. Genome-­wide identification of micro-­ribonucleic acids associated with human endometrial receptivity in natural and stimulated cycles by deep sequencing. Fertil Steril. 2011;96:150–155.e5. 573. May KE, Conduit-­Hulbert SA, Villar J, et al. Peripheral biomarkers of endometriosis: a systematic review. Hum Reprod Update. 2010;16:651–674. 574. May KE, Villar J, Kirtley S, et al. Endometrial alterations in endometriosis: a systematic review of putative biomarkers. Hum Reprod Update. 2011;17:637–653. 575. Lessey BA. Assessment of endometrial receptivity. Fertil Steril. 2013;96:522–529. 576. Ruck P, Marzusch K, Kaiserling E, et al. Distribution of cell adhesion molecules in decidua of early human pregnancy. An immunohistochemical study. Lab Invest. 1994;71:94–101. 577. Lai TH, Zhao Y, Shih IM, et al. Expression of L-­selectin ligands in human endometrium during the implantation window after controlled ovarian stimulation for oocyte donation. Fertil Steril. 2006;85:761–763. 578. Shamonki MI, Kligman I, Shamonki JM, et al. Immunohistochemical expression of endometrial L-­ selectin ligand is higher in donor egg recipients with embryonic implantation. Fertil Steril. 2006;86:1365–1375. 579. Margarit L, Gonzalez D, Lewis PD, et al. L-­selectin ligands in human endometrium: comparison of fertile and infertile subjects. Hum Reprod. 2009;24:2767–2777. 580. Nardo LG, Bartoloni G, Di Mercurio S, et al. Expression of the alpha(v)beta3 and alpha4 beta1 integrins throughout the putative window of implantation in a cohort of healthy fertile women. Acta Obstet Gynecol Scand. 2002;81:753–758. 581. Nardo LG, Nikas G, Makrigiannakis A, et al. Synchronous expression of pinopodes and alpha v beta 3 and alpha 4 beta 1 integrins in the endometrial surface epithelium of normally menstruating women during the implantation window. J Reprod Med.

CHAPTER 10  Structure, Function, and Evaluation of the Female Reproductive Tract253.e13 582. Gonzalez RR, Palomino A, Boric A, et al. A quantitative evaluation of α1, α4, αV and 3 endometrial integrins of fertile and unexplained infertile women during the menstrual cycle. A flow cytometric appraisal. Hum Reprod. 1999;14:2485–2492. 583. Ordi J, Creus M, Ferrer B, et al. Midluteal endometrial biopsy and alphavbeta3 integrin expression in the evaluation of the endometrium in infertility: implications for fecundity. Int J Gynecol Pathol. 2002;21:231–238. 584. Thomas K, Thomson AJ, Sephton V, et al. The effect of gonadotrophic stimulation on integrin expression in the endometrium. Hum Reprod. 2002;17:63–68. 585. Creus M, Balasch J, Ordi J, et al. Integrin expression in normal and out-­of-­phase endometria. Hum Reprod. 1998;13:3460–3468. 586. Daftary G, Troy P, Bagot C, et al. β3 integrin is directly regulated by HOXA10. J Soc Gynecol Investig. 2000;7:55A. 587. Taylor HS, Bagot C, Kardana A, et al. HOX gene expression is altered in the endometrium of women with endometriosis. Hum Reprod. 1999;14:1328–1331. 588. Daftary GS, Taylor HS. Hydrosalpinx fluid diminishes endometrial cell HOXA10 expression. Fertil Steril. 2002;78:577–580. 589. Szczepanska M, Wirstlein P, Luczak M, et al. Reduced expression of HOXA10 in the midluteal endometrium from infertile women with minimal endometriosis. Biomed Pharmacother. 2010;64:697–705. 590. Lessey BA, Castelbaum AJ, Buck CA, et al. Further characterization of endometrial integrins during the menstrual cycle and in pregnancy. Fertil Steril. 1994;62:497–506. 591. Franasiak JM, Holoch KJ, Yuan L, et al. Prospective assessment of midsecretory endometrial leukemia inhibitor factor expression versus alphanubeta3 testing in women with unexplained infertility. Fertil Steril. 2014;101:1724–1731. 592. Carson DD, Lagow E, Thathiah A, et al. Changes in gene expression during the early to mid-­luteal (receptive phase) transition in human endometrium detected by high-­density microarray screening. Mol Hum Reprod. 2002;8:871–879. 593. Talbi S, Hamilton AE, Vo KC, et al. Molecular phenotyping of human endometrium distinguishes menstrual cycle phases and underlying biological processes in normo-­ ovulatory women. Endocrinology. 2006;147:1097–1121. 594. Kao LC, Tulac S, Lobo S, et al. Global gene profiling in human endometrium during the window of implantation. Endocrinology. 2002;143:2119–2138. 595. Riesewijk A, Martin J, van Os R, et al. Gene expression profiling of human endometrial receptivity on days LH+2 versus LH+7 by microarray technology. Mol Hum Reprod. 2003;9:253–264. 596. Koot YE, van Hooff SR, Boomsma CM, et al. An endometrial gene expression signature accurately predicts recurrent implantation failure after IVF. Sci Rep. 2016;6:19411. 597. Ruiz-­Alonso M, Galindo N, Pellicer A, et al. What a difference two days make: “personalized” embryo transfer (pET) paradigm: a case report and pilot study. Hum Reprod. 2014;29:1244–1247. 598. Wang W, Vilella F, Alama P, et al. Single-­cell transcriptomic atlas of the human endometrium during the menstrual cycle. Nat Med. 2020;26(10):1644–1653. 599. Diaz-­Gimeno P, Ruiz-­Alonso M, Blesa D, et al. The accuracy and reproducibility of the endometrial receptivity array is superior to histology as a diagnostic method for endometrial receptivity. Fertil Steril. 2013;99:508–517. 600. Ruiz-­Alonso M, Blesa D, Diaz-­Gimeno P, et al. The endometrial receptivity array for diagnosis and personalized embryo transfer as a treatment for patients with repeated implantation failure. Fertil Steril. 2013;100:818–824. 601. Horcajadas JA, Pellicer A, Simon C. Wide genomic analysis of human endometrial receptivity: new times, new opportunities. Hum Reprod Update. 2007;13:77–86. 602. Garrido-­Gomez T, Ruiz-­Alonso M, Blesa D, et al. Profiling the gene signature of endometrial receptivity: clinical results. Fertil Steril. 2013;99:1078–1085. 603. Gomez E, Ruiz-­Alonso M, Miravet J, et al. Human endometrial transcriptomics: implications for embryonic implantation. Cold Spring Harb Perspect Med. 2015;5:a022996. 604. Garrido-­Gómez T, Ruiz-­ Alonso M, Blesa D, Diaz-­ Gimeno P, Vilella F, Simón C. Profiling the gene signature of endometrial receptivity: clinical results. Fertil Steril. 2013;99(4):1078–1085. 605. Garcia-­Velasco JA, Fassbender A, Ruiz-­ Alonso M, Blesa D, D’Hooghe T, Simon C

affected in women with endometriosis? A pilot study. Reprod Biomed Online. 2015;31(5):647–654. 606. Mahajan N. Endometrial receptivity array: clinical application. J Hum Reprod Sci. 2015;8(3):121–129. 607. Ruiz-­Alonso M, Galindo N, Pellicer A, Simón C. What a difference two days make: “personalized” embryo transfer (pET) paradigm: a case report and pilot study. Hum Reprod. 2014;29(6):1244–1247. 608. Simón C, Gómez C, Cabanillas S, et al. A 5-­year multicentre randomized controlled trial comparing personalized, frozen and fresh blastocyst transfer in IVF. Reprod Biomed Online. 2020;41(3):402–415. 609. Khoufache K, Michaud N, Harir N, et al. Anomalies in the inflammatory response in endometriosis and possible consequences: a review. Minerva Endocrinol. 2012;37:75–92. 610. Weed JC, Arguembourg PC. Endometriosis: can it produce an autoimmune response resulting in infertility? Clin Obstet Gynecol. 1980;23:885. 611. Taylor RN, Kane MA, Sidell N. Pathogenesis of endometriosis: roles of retinoids and inflammatory pathways. Semin Reprod Med. 2015;33:246–256. 612. Li X, Large MJ, Creighton CJ, et al. COUP-­TFII regulates human endometrial stromal genes involved in inflammation. Mol Endocrinol. 2013;27:2041–2054. 613. Fox C, Morin S, Jeong JW, et al. Local and systemic factors and implantation: what is the evidence? Fertil Steril. 2016;105:873–884. 614. Wei Q, Levens ED, Stefansson L, et al. Indian Hedgehog and its targets in human endometrium: menstrual cycle expression and response to CDB-­2914. J Clin Endocrinol Metab. 2010;95:5330–5337. 615. Kurihara I, Lee DK, Petit FG, et al. COUP-­TFII mediates progesterone regulation of uterine implantation by controlling ER activity. PLoS Genet. 2007;3:e102. 616. Lin SC, Li YH, Wu MH, et al. Suppression of COUP-­TFII by proinflammatory cytokines contributes to the pathogenesis of endometriosis. J Clin Endocrinol Metab. 2014;99:E427–E437. 617. Ahn SH, Edwards AK, Singh SS, et al. IL-­ 17A contributes to the pathogenesis of endometriosis by triggering proinflammatory cytokines and angiogenic growth factors. J Immunol. 2015;195:2591–2600. 618. Ahn SH, Khalaj K, Young SL, et al. Immune-­inflammation gene signatures in endometriosis patients. Fertil Steril. 2016;106(6):1420– 1431.e7. 619. Monsanto SP, Edwards AK, Zhou J, et al. Surgical removal of endometriotic lesions alters local and systemic proinflammatory cytokines in endometriosis patients. Fertil Steril. 2016;105:968–977.e5. 620. Copperman AB, Wells V, Luna M, et al. Presence of hydrosalpinx correlated to endometrial inflammatory response in vivo. Fertil Steril. 2006;86:972–976. 621. Kim BG, Yoo JY, Kim TH, et al. Aberrant activation of signal transducer and activator of transcription-­3 (STAT3) signaling in endometriosis. Hum Reprod. 2015;30:1069–1078. 622. Yoo JY, Jeong JW, Fazleabas AT, et al. Protein inhibitor of activated STAT3 (PIAS3) is down-­regulated in eutopic endometrium of women with endometriosis. Biol Reprod. 2016;95(1):11. 623. Evans-­Hoeker E, Lessey BA, Jeong JW, et al. Endometrial BCL6 overexpression in eutopic endometrium of women with endometriosis. Reprod Sci. 2016;23(9):1234–1241. 624. Tiberi L, Bonnefont J, van den Ameele J, et al. A BCL6/BCOR/SIRT1 complex triggers neurogenesis and suppresses medulloblastoma by repressing Sonic Hedgehog signaling. Cancer Cell. 2014;26:797–812. 625. Bukulmez O, Hardy DB, Carr BR, et al. Inflammatory status influences aromatase and steroid receptor expression in endometriosis. Endocrinology. 2008;149:1190–1204. 626. Tamai K, Togashi K, Ito T, et al. MR imaging findings of adenomyosis: correlation with histopathologic features and diagnostic pitfalls. Radiographics. 2005;25:21–40. 627. Ferenczy A. Pathophysiology of adenomyosis. Hum Reprod Update. 1998;4:312–322. 628. Vercellini P, Vigano P, Somigliana E, et al. Adenomyosis: epidemiological factors. Best Pract Res Clin Obstet Gynaecol. 2006;20:465–477. 629. Campo S, Campo V, Benagiano G. Adenomyosis and infertility. Reprod Biomed Online. 2012;24:35–46. 630. Harada T, Khine YM, Kaponis A, et al. The impact of adenomyosis on women’s fertility. Obstet Gynecol Surv. 2016;71:557–568. 631. Abbott JA. Adenomyosis and abnormal uterine bleeding (AUB-­A)-­ pathogenesis, diagnosis, and management. Best Pract Res Clin Obstet

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253.e14 PART I  The Fundamentals of Reproduction 632. Oh SJ, Shin JH, Kim TH, et al. beta-­Catenin activation contributes to the pathogenesis of adenomyosis through epithelial-­mesenchymal transition. J Pathol. 2013;231:210–222. 633. Parazzini F, Vercellini P, Panazza S, et al. Risk factors for adenomyosis. Hum Reprod. 1997;12:1275–1279. 634. Vavilis D, Agorastos T, Tzafetas J, et al. Adenomyosis at hysterectomy: prevalence and relationship to operative findings and reproductive and menstrual factors. Clin Exp Obstet Gynecol. 1997;24:36–38. 635. Pontis A, D’Alterio MN, Pirarba S, et al. Adenomyosis: a systematic review of medical treatment. Gynecol Endocrinol. 2016;32:696–700. 636. Bartel DP. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell. 2004;116:281–297. 637. Cai X, Hagedorn CH, Cullen BR. Human microRNAs are processed from capped, polyadenylated transcripts that can also function as mRNAs. RNA. 2004;10:1957–1966. 638. Cullen BR. Derivation and function of small interfering RNAs and microRNAs. Virus Res. 2004;102:3–9. 639. Toloubeydokhti T, Pan Q, Luo X, et al. The expression and ovarian steroid regulation of endometrial micro-­ RNAs. Reprod Sci. 2008;15:993–1001. 640. Pan Q, Chegini N. MicroRNA signature and regulatory functions in the endometrium during normal and disease states. Semin Reprod Med. 2008;26:479–493. 641. Pan Q, Luo X, Toloubeydokhti T, et al. The expression profile of micro-­RNA in endometrium and endometriosis and the influence of ovarian steroids on their expression. Mol Hum Reprod. 2007;13:797–806. 642. Aghajanova L, Giudice LC. Molecular evidence for differences in endometrium in severe versus mild endometriosis. Reprod Sci. 2011;18:229–251. 643. Ramon LA, Braza-­Boils A, Gilabert-­Estelles J, et al. microRNAs expression in endometriosis and their relation to angiogenic factors. Hum Reprod. 2011;26:1082–1090. 644. Grechukhina O, Petracco R, Popkhadze S, et al. A polymorphism in a let-­7 microRNA binding site of KRAS in women with endometriosis. EMBO Mol Med. 2012;4:206–217. 645. Cosar E, Mamillapalli R, Ersoy GS, et al. Serum microRNAs as diagnostic markers of endometriosis: a comprehensive array-­based analysis. Fertil Steril. 2016;106:402–409. 646. Nisenblat V, Bossuyt PM, Shaikh R, et al. Blood biomarkers for the non-­invasive diagnosis of endometriosis. Cochrane Database Syst Rev. 2016;(5):CD012179. 647. Okamoto M, Nasu K, Abe W, et al. Enhanced miR-­210 expression promotes the pathogenesis of endometriosis through activation of signal transducer and activator of transcription 3. Hum Reprod. 2015;30:632–641. 648. Zhou M, Fu J, Xiao L, et al. miR-­196a overexpression activates the MEK/ERK signal and represses the progesterone receptor and decidualization in eutopic endometrium from women with endometriosis. Hum Reprod. 2016;31(11):2598–2608. 649. Liu XJ, Bai XG, Teng YL, et al. miRNA-­ 15a-­ 5p regulates VEGFA in endometrial mesenchymal stem cells and contributes to the pathogenesis of endometriosis. Eur Rev Med Pharmacol Sci. 2016;20:3319–3326. 650. Taylor E, Gomel V. The uterus and fertility. Fertil Steril. 2008;89:1–16. 651. Amso NN, Griffiths A. The role and applications of ultrasound in ambulatory gynaecology. Best Pract Res Clin Obstet Gynaecol. 2005;19:693–711. 652. Bassil S. Changes in endometrial thickness, width, length and pattern in predicting pregnancy outcome during ovarian stimulation in in vitro fertilization. Ultrasound Obstet Gynecol. 2001;18:258–263.

653. Gonen Y, Calderon I, Dirnfeld M, et al. The impact of sonographic assessment of the endometrium and meticulous hormonal monitoring during natural cycles in patients with failed donor artificial insemination. Ultrasound Obstet Gynecol. 1991;1:122–126. 654. Ohno Y, Hosokawa K, Tamura T, et al. Endometrial oestrogen and progesterone receptors and their relationship to sonographic endometrial appearance. Hum Reprod. 1995;10:708–711. 655. Puerto B, Creus M, Carmona F, et al. Ultrasonography as a predictor of embryo implantation after in vitro fertilization: a controlled study. Fertil Steril. 2003;79:1015–1022. 656. Sterzik K, Grab D, Schneider V, et al. Lack of correlation between ultrasonography and histologic staging of the endometrium in in vitro fertilization (IVF) patients. Ultrasound Med Biol. 1997;23:165–170. 657. Battaglia C, Sgarbi L, Salvatori M, et al. Increased anticardiolipin antibodies are positively related to the uterine artery pulsatility index in unexplained infertility. Hum Reprod. 1998;13:3487–3491. 658. Habara T, Nakatsuka M, Konishi H, et al. Elevated blood flow resistance in uterine arteries of women with unexplained recurrent pregnancy loss. Hum Reprod. 2002;17:190–194. 659. Carbillon L, Perrot N, Uzan M, et al. Doppler ultrasonography and implantation: a critical review. Fetal Diagn Ther. 2001;16:327–332. 660. Goldstein SR. Controversy about uterine effects and safety of SERMs: the saga continues. Menopause. 2002;9:381–384. 661. Maymon R, Herman A, Ariely S, et al. Three-­dimensional vaginal sonography in obstetrics and gynaecology. Hum Reprod Update. 2000;6:475–484. 662. Alcazar JL. Three-­dimensional ultrasound assessment of endometrial receptivity: a review. Reprod Biol Endocrinol. 2006;4:56. 663. van Dongen H, de Kroon CD, Jacobi CE, et al. Diagnostic hysteroscopy in abnormal uterine bleeding: a systematic review and meta-­ analysis. BJOG. 2007;114:664–675. 664. Jansen FW, Vredevoogd CB, van Ulzen K, et al. Complications of hysteroscopy: a prospective, multicenter study. Obstet Gynecol. 2000;96:266–270. 665. Phillips K. Hysteroscopic sterilisation: is laparoscopic sterilisation a thing of the past? BJOG. 2007;114:661–663. 666. James AH, Kouides PA, Abdul-­Kadir R, et al. Von Willebrand disease and other bleeding disorders in women: consensus on diagnosis and management from an international expert panel. Am J Obstet Gynecol. 2009;201:12.e1–12.e8. 667. ACOG. Management of abnormal uterine bleeding associated with ovulatory dysfunction. ACOG Practice bulletin no. 136. Obstet Gynecol. 2013;122:176–185. 668. Abbott DH, Dumesic DA, Eisner JR, et al. Insights into the development of polycystic ovary syndrome (PCOS) from studies of prenatally androgenized female rhesus monkeys. Trends Endocrinol Metab. 1998;9:62–67. 669. Carmina E, Lobo RA. Polycystic ovary syndrome (PCOS): arguably the most common endocrinopathy is associated with significant morbidity in women. J Clin Endocrinol Metab. 1999;84:1897–1899. 670. Rotterdam ESHRE/ASRM-­Sponsored PCOS Consensus Workshop Group. Revised 2003 consensus on diagnostic criteria and long-­term health risks related to polycystic ovary syndrome (PCOS). Hum Reprod. 2004;19:41–47. 671. Giudice LC. Endometrium in PCOS: implications and predisposition for endocrine CA. Best Pract Res Clin Endocrinol Metab. 2006;20:235–244. 672. Hickey M, d’Arcangues C. Mechanisms Underlying Menstrual Bleeding Disturbances with Progestogens. Ernst Schering Res Found Workshop; 2005:191–217. 673. Curtis KM, Hillis SD, Marchbanks PA, et al. Disruption of the endometrial-­myometrial border during pregnancy as a risk factor for adenomyosis. Am J Obstet Gynecol. 2002;187:543–544.

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Endocrinology of Human Pregnancy and Fetal-­Placental Neuroendocrine Development Sam Mesiano

OUTLINE INTRODUCTION ESTABLISHMENT OF PREGNANCY Endometrial Receptivity Decidualization of the Endometrial Stroma Implantation Placentation Immune Tolerance of the Conceptus THE PLACENTA AS AN ENDOCRINE ORGAN Chorionic Gonadotropin Gonadotropin-­Releasing Hormone Activins and Inhibins Thyrotropin-­Releasing Hormone Placental Somatotropins Placental Extracellular Vesicles Cell-­Free Fetal DNA FETAL NEUROENDOCRINE SYSTEMS Hypothalamic Hormones Pituitary Hormones Fetal Pituitary-­Adrenal Axis Fetal Pituitary-­Gonadal Axis Fetal Pituitary-­Thyroid Axis Fetal Pituitary-­Growth Hormone Axis Fetal Pituitary-­Prolactin Axis FETAL MATURATION AND THE TIMING OF PARTURITION Fetal Organ Maturation and Preparation for Extrauterine Life Process of Human Parturition Hormonal Control of Human Parturition Fetal Lung Maturation UTERINE STRETCH EVOLUTIONARY PERSPECTIVE

INTRODUCTION The birth of a healthy baby is dependent on an ordered sequence of biological events during pregnancy. Important among these are:    1. Successful implantation of the developing embryo 2. Adaptation of maternal physiology to accept and retain the embryo and satisfy its nutritional, metabolic, and physical needs 3. Appropriate growth and functional development of key organ and homeostatic control systems in the fetus in preparation for life outside the uterus 4. Proper timing of birth so that it occurs when the fetus is mature enough to survive as a neonate   

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The hormonal interactions between the fetus/placenta and mother that control these processes are discussed in this chapter. The chapter begins by considering the hormonal events that prepare the uterus for pregnancy. The uterine endometrium is conditioned during the menstrual cycle by hormones that affect its growth, morphology, and function such that it becomes receptive to embryo implantation. Implantation of the conceptus into the endometrium involves a complex tissue-­level paracrine dialogue involving trophoblast cells of the embryo and cells of the maternal endometrium, its vasculature, and resident immune cells. The process leads to maternal immune tolerance of the conceptus and development of the placenta through which the embryo/fetus obtains nutrients and oxygen from the maternal compartment. This process is critical for the success of pregnancy and is central to the etiology of adverse conditions such as preeclampsia and intrauterine growth restriction. Once pregnancy is established, the fetus, placenta, and mother initiate and maintain communication mainly via the endocrine system. In women, the maternal endocrine milieu of pregnancy is dominated by placental hormones, the major function of which is to modify maternal physiology to maintain pregnancy and satisfy the nutritional, metabolic, and physical needs of the growing fetus. Pregnancy ends with the birth of the fetus and placenta through the process of parturition. The timing of parturition is critical for the success of pregnancy. Birth normally occurs when the fetus is sufficiently mature to survive as a newborn. To this end, functional maturation of the neuroendocrine system must occur in the fetus so that it is ready for life as a newborn. The development of key neuroendocrine axes in the human fetus is discussed. Hormonal signaling networks between the fetus, placenta, and mother, controlling the process and timing of human parturition, are also considered.

ESTABLISHMENT OF PREGNANCY • Pregnancy involves retention and gestation of the embryo in the uterus. • The establishment of pregnancy requires an endometrium that is permissive to embryo implantation and the development of a placenta. • Paracrine signaling between the embryo and cells in the endometrium is necessary for implantation and placentation. • Endocrine signals from the embryo maintain progesterone production by the corpus luteum to prevent menstruation and allow for the establishment of pregnancy.

Endometrial Receptivity The ovarian steroid hormones, estradiol, and progesterone, prepare the uterus for pregnancy by inducing structural and functional changes in the endometrium (see Chapter 10). The endometrium has two functional layers, the stratum functionalis (closest to the uterine cavity) and the stratum basalis (adjacent to the myometrium). The stratum functionalis is responsive to estradiol and progesterone and forms a proliferative phenotype

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

in response to estradiol or a secretory/decidualized phenotype in response to estradiol and progesterone. The secretory/decidualized stratum functionalis is shed at menstruation, miscarriage, and parturition. In the following cycle, estrogen induces the development of a new stratum functionalis from cells in the stratum basalis. Pregnancy is established in the postovulatory secretory/decidualized endometrium during the luteal phase of the menstrual cycle. Receptivity of the endometrium to embryo adhesion and implantation has temporal and spatial characteristics.1–4 In humans, peak endometrium receptivity occurs between days 21 and 24 of the menstrual cycle. This is referred to as the implantation window. During this time, the probability of successful implantation of a healthy embryo is ∼85%. In contrast, implantation before or after the implantation window has a low probability (∼11%) of establishing a successful pregnancy.5 Analysis of the human endometrial transcriptome during the implantation window has revealed numerous genes that are up-­and downregulated in comparison with late proliferative phase endometrium.6,7 Under the influence primarily of progesterone, the postovulatory endometrium transforms to the secretory phenotype. It thickens and becomes highly vascularized, and the glandular epithelium secretes glycoproteins and other factors to produce an intrauterine environment that favors survival of the floating embryo. The luminal epithelial cells also produce chemokines, growth factors, and cell adhesion molecules (CAMs) that attract the embryo to specific dome-­ shaped docking sites known as pinopodes.8 The tips of the pinopodes express chemokines and CAMs that attract the embryo and appear to be the preferred site for embryo adherence. Areas between pinopodes produce a repellent molecule, MUC-­1, that prevents embryo adhesion.9 Embryo adhesion initiates the invasion of trophoblasts across the epithelium. The interaction also triggers the process of decidualization in the underlying endometrial stroma that is necessary for subsequent implantation and placentation.

Decidualization of the Endometrial Stroma Late in the luteal phase, the endometrial stroma undergoes a morphologic and functional transformation referred to as decidualization.4,10–12 Stromal fibroblasts in the endometrium proliferate and differentiate into large polyhedral epithelioid cells with high levels of glycogen and lipids. The cells also produce a tough pericellular capsule composed of collagen, laminin, fibronectin, and heparan sulfate proteoglycans. In a nonfertile cycle, decidualization begins around blood vessels during the midsecretory phase and progressively encompasses the entire endometrial stroma during the latter part of the secretory phase. In a fertile cycle, decidualization is also triggered by signals from the luminal epithelium in response to blastocyst adhesion. Decidualized endometrial stromal cells communicate with adjacent epithelial cells, immune cells, vascular endothelial cells, and trophoblast cells of the invading embryo to produce a microenvironment that is conducive to the establishment and continuation of pregnancy. Decidual cells promote the growth and remodeling of uterine spiral arterioles and regulate the local immune cell ecosystem to produce an immune-­privileged tolerogenic site conducive to implantation of the semiallogeneic embryo. The specific cohort of resident immune cells in the decidua also restricts the passage of fetal antigens to the maternal compartment to further protect the conceptus from attack by the maternal immune system.13,14 The decidual microenvironment also promotes the development of robust uterine arteriolar vasculature, providing the developing placenta access to the maternal circulation.15 Although decidualized stromal cells surround the implant ing blastocyst and are essential for successful implantation, the

255

fully decidualized endometrium limits the depth of embryo invasion into the uterine wall and is hostile to de novo implantation. Placenta accreta is a condition characterized by excessive and sometimes life-­threatening trophoblast invasion and is thought to be caused by invasion of the embryo into uterine scar tissue (usually from a previous Cesarean section) that lacks decidua.16 Decidualization is dependent on progesterone produced by the corpus luteum (CL) and cyclic adenosine monophosphate (cAMP) signaling in endometrial stromal cells. In decidualized endometrial stromal cells, progesterone and cAMP induce production of multiple regulatory factors, especially prolactin, insulin-­like growth factor (IGF)-­binding protein 1 (IGFBP1), and cytokines such as interleukin (IL)-­15 (IL-­15) that participate in a complex cross-­talk between endometrial epithelial cells and resident immune cells to create a microenvironment conducive for blastocyst implantation. Prolactin and IGFBP1 are thought to promote implantation by stimulating the proliferation and invasive properties of trophoblasts.17,18 IL-­15 promotes the differentiation of resident immune cells, especially uterine natural killer (uNK) cells, to produce an immune microenvironment that is tolerant of the semiallogeneic conceptus. In a nonfertile cycle, the decidualized stratum functionalis is shed at menstruation, in response to progesterone withdrawal due to CL regression. In a fertile cycle, chorionic gonadotropin (CG) produced by the embryo prevents CL regression, leading to sustained progesterone support for the decidualized endometrium, which is necessary for the establishment and maintenance of pregnancy.

Implantation After syngamy, an intrinsic development program is initiated in the early embryo (see Chapter 9). The sequence of cell divisions and differentiation is not dependent on the hormonal milieu of the fallopian tube or the uterus. By the third to fourth day after fertilization, the embryo comprises a solid ball of cells encapsulated by the translucent remnant of the zona pellucida. On about the fifth day after fertilization, a fluid-­filled cavity, the blastocele, forms within the embryo, which at this stage is referred to as a blastocyst. The outer layer of blastocyst cells adjacent to the zona pellucida is known as the trophectoderm. These cells directly interact with maternal tissue and eventually give rise to the placenta and chorion. The fetus and the amnion and mesenchymal and vascular components of the placenta are derived from a group of cells, known as the inner cell mass, lying under the trophectoderm at one end of the blastocele. The human blastocyst enters the uterus around the fifth day after fertilization and floats freely for 2 to 3 days. By this stage, the blastocyst escapes the confines of the zona pellucida (i.e., it hatches) and is in an optimal condition to implant into a receptive endometrium. Almost immediately after fertilization, hormonal signals from the conceptus are transmitted to the mother. Studies of the secreted proteins (the secretome) produced by the preimplantation embryo show that the profile of secreted proteins changes every 24 hours and with different embryonic stages.20–22 This suggests that the embryo communicates with the uterine epithelium via a specifically orchestrated sequence of secretions. Any delay in embryo development or its transit through the oviduct increases the likelihood of implantation failure. Given this delicate balance, it is not surprising that the highest rate of pregnancy loss occurs during the periimplantation period. The embryo develops and implants in a polarized fashion. After hatching from the zona pellucida, trophoblast cells overlying the inner cell mass of the embryonic pole are the first to interact with the uterine epithelium, most likely at the

11

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Placentation

Uterine stroma Invading syncytiotrophoblast Trophoblast

Uterine epithelium Inner cell mass (embryonic disk) Cytotrophoblast Blastocele

A

B Uterine stroma Syncytiotrophoblast Amnionic cavity Endoderm Cytotrophoblast Blastocele (yolk sac cavity)

C Sinusoids filled with maternal blood Amnionic cavity Syncytiotrophoblast Yolk sac cavity

D Fig. 11.1 Implantation of the human embryo. (A) Floating blastocyst. (B) Attachment to the uterine epithelium and initial invasion of the syncytiotrophoblast cells. (C) The blastocyst penetrates deeper into the uterine stroma and develops an amniotic cavity. (D) The fully implanted embryo invades the maternal vasculature and the uterine epithelium grows over the implantation site and undergoes decidualization. (From Jones RE, ed. Human Reproductive Biology. Academic Press; 1997:189.)

pinopodes8,23–25 (Fig. 11.1). The trophoblast cells proliferate and secrete proteases that degrade the extracellular matrix between endometrial cells, forming a path for the blastocyst to enter the uterine stroma. At this stage, the trophoblast cells are referred to as cytotrophoblasts and differentiate along two distinct paths: villous and extravillous. The extravillous cytotrophoblasts become highly invasive and form columns penetrating the basal membrane beneath the endometrial stroma and into the myometrium. Eventually, the entire embryo is embedded in the uterine stroma and anchored by columns of extravillous cytotrophoblasts. During this time, some villous cytotrophoblasts fuse their plasma membranes to become a single multinucleated cell known as the syncytiotrophoblast, which becomes the outer layer of the placental villi (see below). As with adhesion, implantation also involves a paracrine dialogue between the embryo and the endometrial stoma. Some of the molecules involved in the paracrinology of implantation include3,26–30:    • Leukemia inhibitory factor31 • Interleukin-­1132 • Heparin-­binding epidermal growth factor-­like growth factor33 • Prostaglandins, especially prostacyclin and prostaglandin E234 • The homeobox transcription factors, especially HOXA10 and HOXA1135,36 • Metalloproteinases37 • The Wnt and Indian

Early in the implantation process some cytotrophoblast cells aggregate and form migrating columns that penetrate into the inner third of the myometrium. The invading columns of cytotrophoblasts target maternal blood vessels and, via interstitial and endovascular routes, completely surround and occlude spiral arterioles. They then displace maternal endothelial cells and vascular smooth muscle cells to create a low-­resistance arteriolar system by increasing vessel diameter. Poiseuille’s law of fluid dynamics dictates that flow through a cylinder is proportional to its radius multiplied by the fourth power. As a consequence of cytotrophoblast invasion, the average radius of human uterine arterioles is around 10-­fold greater during pregnancy compared with the nonpregnant state. Such a change increases flow 10,000-­ fold. Cytotrophoblasts within the spiral arterioles also prevent maternal vasomotor control of the spiral arterioles. Through this process, the supply of nutrients and oxygen to the conceptus is optimized and the capacity for maternal restriction of uterine blood flow is subverted. Maternal blood flowing from the dilated spiral arterioles fills spaces, known as lacunae, that form between the invading columns of cytotrophoblast cells. At around the same stage of placental development, finger-­like projections of chorionic villi form in the lacunae and become bathed in maternal blood. Chorionic villi have a central core of loose connective tissue with an extensive capillary network linked with fetal circulation. Surrounding the cores are the outer syncytiotrophoblast and inner cytotrophoblast cells that form a barrier between the maternal and fetal circulations. This arrangement is referred to as hemochorial placentation since the maternal blood is in direct contact with the syncytiotrophoblast and extravillous cytotrophoblasts. Placentation across all eutherian mammals is characterized by high angiogenic activity and blood vessel growth,40 especially at the site of placental attachment. The cytotrophoblast cells produce several angiogenic factors, including platelet-­derived endothelial cell growth factor, vascular endothelial growth factor (VEGF), angiopoietin-­1, and angiopoietin-­2.41,42 In addition, two potent inhibitors of angiogenesis have been isolated from mouse placenta.43 The presence of antiangiogenic factors during placentation is thought to prevent maternal endothelial cells from resealing the ends of spiral arterioles that have been occupied by cytotrophoblasts. In addition, antiangiogenic factors may prevent the overgrowth of maternal and fetal vessels, thereby preventing maternal blood vessels from entering the fetal compartment and fetal vessels from extending beyond the uterus. Angiogenic events occurring during implantation and placentation are thought to be critical factors in the etiology of hemodynamic disorders in pregnancy. The extent and depth of the placental incursion appear to be the sum of the intrinsic proinvasive characteristics of cytotrophoblasts and the physical and biochemical barriers mounted by the maternal tissues. Imbalances in this equation can lead to failed implantation and pathological conditions. The arteriole remodeling process is complex and requires the cytotrophoblast cells to express specific adhesion molecules and adopt an endothelial phenotype. If this process is impaired the depth of invasion and remodeling may be limited, leading to placental ischemia (Fig. 11.2). Abnormal spiral artery remodeling is associated with fetal growth restriction and hypertensive disorders of pregnancy.44,45 In preeclampsia, cytotrophoblast invasion is restricted to the superficial decidual segments, leaving the myometrial spiral arterioles undisturbed and still responsive to maternal vasomotor control.46 To compensate, especially during the later stages of pregnancy when nutritional requirements of the fetus increase, the placenta is thought to secrete factors into the maternal circulation that increase maternal systemic

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

257

Normal Syncytiotrophoblast

11 Myometrium

Deoxygen

ated blood

Maternal vein Cytotrophoblast stem cells

Fetal blood vessels Cytotrophoblast Tunica Media smooth muscle

Maternal Blood

Anchoring villus

Blood

Flow

Spiral Artery

Decidua Floating villus

Anchoring villus cytotrophoblast column

Cytotrophoblast

Maternal endothelial cells

Endovascular cytotrophoblast

Preeclampsia Syncytiotrophoblast

Myometrium

Deoxygen ate

d blood

Maternal vein Cytotrophoblast stem cells

Fetal blood vessels Cytotrophoblast

Tunica Media smooth muscle

Maternal Blood

Anchoring villus

Bloo d Flo w

Spiral Artery

Floating villus

Anchoring villus cytotrophoblast column

Decidua

Maternal endothelial cells

Fig. 11.2 Invasion of cytotrophoblast cells into the spiral arteries. Shallow invasion restricts blood flow to the placenta and is thought to be a precursor for the development of preeclampsia. (From Lam C, Lim KH, Karumanchi SA. Circulating angiogenic factors in the pathogenesis and prediction of preeclampsia. Hypertension. 2005;46(5):1077–1085.)

Immune Tolerance of the Conceptus Because cytotrophoblasts are genetically and immunologically distinct from maternal tissue, implantation represents an extraordinary breach of maternal immune defenses. In 1953, Sir Peter Medawar in his studies of graft rejection proposed 3 mechanisms by which the embryo is protected from maternal immune rejection: (1) anatomical separation between fetal cells and maternal immune cells; (2) modulation of the maternal immune cells at the maternal-­fetal interface; and/or (3) global modulation of the maternal immune system to produce refractoriness to allogenic fetal cells.47 It is now clear that a physical barrier between maternal immune cells and fetal cells expressing alloantigens does not exist,

thus eliminating Medawar’s first mechanism. Medawar’s second and third mechanisms are supported by clinical and animal data. Although maternal lymphocytes with the capacity to attack and eliminate fetal cells exist throughout gestation,48,49 they are suppressed during pregnancy.50 This tolerogenic state is exemplified clinically by reduced severity of autoimmune disease during pregnancy and its relapses after parturition,51 and experimentally by increased tolerance for engraftment of paternally derived tumor cells in sites outside of the uterus during pregnancy in mice.48 The tolerogenic state of pregnancy is thought to involve a subset of T cells known as regulatory T (T-­reg) cells that promote tolerance to nonself antigens, inhibit inappropriate immune responses against self-antigen, and whose population increases during pregreg cell expansion is synergistically

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TABLE 11.1  Hormones Produced by the Human Placenta Neuropeptides

Pituitary-­Like Hormones

Adipokine

Growth Factors

Steroid Hormones

CRH TRH GnRH Melatonin Cholecystokinin Met-­enkephalin Dynorphin Neurotensin VIP Galanin Somatostatin CGRP Neuropeptide Y Substance P Endothelin ANP Renin Angiotensin Urocortin Kisspeptin

ACTH TSH PGH PL CG LH FSH β-­Endorphin Prolactin Oxytocin Activin Follistatin Inhibin

Adiponectin Leptin Resistin Visfatin Ghrelin FGF21

IGF-­I/-­II VEGF EGF

Progesterone Estradiol Estrone Estriol Estetrol 2-­Methoxyestradiol Allopregnanolone Pregnenolone 5α-­Dihydroprogesterone

Monoamines and Adrenal-­Like Peptides Epinephrine Norepinephrine Dopamine Serotonin Adrenomedullin

ANP, Atrial natriuretic peptide; CGRP, calcitonin gene-­related peptide; FGF21, fibroblast growth factor 21; PAPP-­A, pregnancy-­associated plasma protein A; VIP, vasoactive intestinal peptide. (Adapted from Reis FM, Petraglia F. The placenta as a neuroendocrine organ. Front Horm Res. 2001;27:216 and Costa MA. The endocrine function of the human placenta: an overview. Reprod BioMed Online. 2016;32:14–43.)

driven by exposure to fetal alloantigen55 and by the endocrine milieu of pregnancy, especially by progesterone.56 Progesterone also stimulates Fas-­ ligand production by decidual cells that induce apoptosis of activated maternal T lymphocytes.57–59 Progesterone also stimulates decidual stromal cells to produce cytokines, especially IL-­15, that induce uNKs with immunosuppressive activity.60,61 Cytotrophoblast cells avoid detection by maternal immune cells by producing a distinct human leukocyte antigens (HLA) profile involving polymorphic molecules, especially HLA-­G, that do not trigger NK cell activation.62–66 Cytotrophoblast cells also may inhibit the proliferation of maternal T cells by catabolizing tryptophan, an essential amino acid needed for proliferating T cells.67,68

THE PLACENTA AS AN ENDOCRINE ORGAN The endocrine milieu of pregnancy is dominated by the placenta. • • The human placenta has direct access to the maternal circulation. • Placental cells produce hormones that modulate maternal physiology to maintain pregnancy and provide the fetus with resources needed for growth and development.    Placental hormones dominate the endocrine milieu of human pregnancy.69,70 This remarkable organ produces a plethora of hormones that it secretes in large quantities into the maternal circulation (Table 11.1). The hemochorial arrangement of the human placenta is ideal for this purpose with the syncytiotrophoblast of the chorionic villi having direct access to maternal blood. Most placental hormones are identical to those produced in the nonpregnant adult and therefore bind to the same cognate receptors. In this context, placental hormones may be regarded as allocrine factors, that is, hormones produced by one organism, the fetus, to act on the receptors of another, the mother. Remarkably, the placenta produces and responds to hypothalamic/pituitary hormone analogs and as such functions as a secondary neuroendocrine control center.70,71 This “extra brain” in many instances overrides maternal systems to affect maternal physiology, usually in favor of pregnancy and provisioning the conceptus with nutrients and oxygen. The role of some of the

principal placental hormones in the endocrine control of human pregnancy is discussed below.

Chorionic Gonadotropin One of the first endocrine signals from the embryo to the mother is mediated by a gonadotropin-­like glycoprotein hormone, chorionic gonadotropin (CG; also known as hCG in humans) (see Chapter 2). CG is produced by trophoblast cells of the early embryo and binds to luteinizing hormone (LH) receptors on CL cells, where it acts as a super-­gonadotropin to prolong the longevity and steroidogenic function of the CL. Importantly, CG prevents regression of the CL that would otherwise occur in a nonfertile cycle and as such sustains progesterone synthesis required to prevent menstruation and establish pregnancy. This is referred to clinically as the maternal recognition of pregnancy. CG is biologically similar to pituitary LH and structurally similar to LH and follicle-­stimulating hormone (FSH). It is a heterodimer composed of α and β subunits. The α-­subunit of CG, LH, and FSH are identical, whereas the β-­subunits differ. The α subunit of CG is produced by cytotrophoblast cells especially as they differentiate during implantation. The β subunit of CG is primarily produced by the syncytiotrophoblast but can also be detected in mature cytotrophoblast cells just before they fuse to form the syncytiotrophoblast. The syncytiotrophoblast produces both subunits and is the principal source of CG. As a potent luteotropin, CG acts via the LH receptor and stimulates cells of the CL to produce and release progesterone. In normal pregnancies, CG is detectable (by measuring β-­ CG) 9 to 11 days after the midcycle LH peak, which is around 8 days after ovulation and only 1 day after implantation. This has clinical utility when it is important to determine the presence of pregnancy at an early stage. In early pregnancy, circulating CG levels double every 2 to 3 days, and concentrations of CG rise to peak values by 60 to 90 days of gestation. Thereafter, CG levels decrease to a plateau that is maintained during the remainder of the pregnancy (Fig. 11.3). Assayable LH and FSH levels in the maternal blood are virtually undetectable throughout pregnancy. Actions of CG may not be limited to maintaining progesterMuch of the increased maternal

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

Serum level (µg/mL)

8

hPL hCG

6

4

2

0 0

10

20

30

40

Weeks from LMP Fig. 11.3 Schematic representation of concentrations of human chorionic gonadotropin (CG) and placental lactogen (PL) throughout gestation. Note differences in the magnitude of the concentrations of the two hormones in early and late gestation. LMP, Last menstrual period.

thyroid activity that occurs in pregnancy has been attributed to CG,73 which binds specifically to thyroid cell membranes and displaces thyroid-­stimulating hormone (TSH).74 CG also influences the development and function of the fetal adrenals and testes.75 In addition, CG may have actions on the maternal reproductive tract, including the decidual response,76 relaxin production by the CL,77 and relaxation of uterine smooth muscle.78,79 LH receptors have been detected in fetal membranes.80 Extravillous cytotrophoblasts also produce hyperglycosylated CG and monomeric β-­subunit that also act via the LH receptor to affect other tissues including some malignancies.72,81–83 Production of CG by the invading blastocyst may contribute in a paracrine manner to the implantation process. In vitro and in vivo studies in nonhuman primates demonstrate that CG promotes decidualization of the endometrial stroma.84 The hormone affects endometrial cells at the implantation site before its levels are detectable in the circulation. Studies suggest that CG produced by the blastocyst prolongs the window of implantation by inhibiting endometrial IGFBP1 production, augmenting angiogenesis at the implantation site by increasing VEGF expression, modulating local cytokine and chemokine expression, augmenting local protease activity and, via an autocrine effect on cytotrophoblasts, augmenting the invasive potential of the blastocyst.72,76

Gonadotropin-­Releasing Hormone The human placenta produces GnRH, which is identical to that produced by the hypothalamus.85 Levels of GnRH in the circulation of pregnant women are highest in the first trimester and correlate closely with CG levels.86 The close relation between GnRH and CG suggests a role for GnRH in regulating CG production.87 GnRH stimulates the production of both the α-­subunit and β-­subunit of CG in placental explants and specific GnRH-­ binding sites are present in the human placenta.85 Other regulators of pituitary gonadotropin, such as progesterone, inhibin, and activin, also influence placental CG production (at least in vitro).

Activins and Inhibins Activin and inhibin are disulfide-­linked homo-­and heterodimeric proteins belonging to the transforming growth factor -­β (TGF-­β) superfamily. Inhibin is a heterodimer composed of an α subunit and one of two

259

inhibit pituitary FSH secretion. In contrast, activins, composed of βA or βB homodimers (βA-­βA and βB-­βB), stimulate FSH production. Both hormones affect target cell function via specific cell surface receptors. The bioavailability of activin is restricted by follistatin, which binds to activin and prevents it from interacting with its receptor on target cells. Inhibins and activins are produced by the human placenta. Each of the subunits is expressed in the syncytiotrophoblast and the levels of expression do not change with advancing gestation.88–90 Activin-­A is also produced by the CL, decidua, and fetal membranes.91 The placenta also produces follistatin.92 These factors are secreted into the maternal and fetal circulations and amniotic fluid and their production varies with the stage of gestation.93 Although the exact function of the inhibin-­activin system in human pregnancy is not known, several studies indicate their involvement, and those of other TGF-­ β family members, in placental development and function, and in the pathogenesis of adverse pregnancy conditions.94 Levels of inhibin-­A and activin­A in the maternal circulation can be indicative, albeit with relatively weak predictive value, of pathologies such as placental tumors, hypertensive disorders of pregnancy, intrauterine growth restriction, fetal hypoxia, Down syndrome, fetal demise, preterm delivery, and intrauterine growth restriction.95–98 Because inhibin and activin are synthesized by cytotrophoblast cells, they may be involved in autoregulating placental CG production by modulating local GnRH activity. In vitro studies have shown that inhibin decreases GnRH-­stimulated CG production by placental cell cultures, whereas antibody blockade of inhibin activity increases GnRH release and causes a parallel rise in CG secretion.99 Thus, it is possible that inhibin exerts an autocrine effect on placental CG secretion by suppressing GnRH action. In contrast, activin augments the GnRH-­induced release of CG in cultured trophoblast cells, an effect that can be reduced by the addition of inhibin.99 Thus, at least in vitro, activin and inhibin contribute to the regulation of CG secretion in a manner similar to their effect on hypothalamic-­pituitary gonadotropin secretion.100

Corticotrophin-­Releasing Hormone First identified in the hypothalamus, corticotrophin-­releasing hormone (CRH) is a 41-­amino acid peptide that stimulates the expression and processing of proopiomelanocortin (POMC) by pituitary corticotropes, and the secretion of adrenocorticotropic hormone (ACTH), a key POMC derivative. The human placenta, fetal membranes, and decidua also produce CRH that is identical to that produced by the hypothalamus.101 CRH produced by the placenta can be detected from the seventh week of pregnancy and increases progressively until term, rising more than 20-­fold in the last 5 to 7 weeks of pregnancy.102 Placental CRH is released mainly into the maternal compartment. Levels of CRH in the maternal circulation can be detected as early as 15 weeks of gestation and then increase through gestation, reaching maximum levels of 1 to 10 ng/mL at term. Remarkably, this is about 1000-­fold higher than peripheral CRH levels in nonpregnant women.103 Actions of CRH are mediated by two CRH receptors (CRH-­Rs), CRH-­R1 and CRH-­R2.104,105 A CRH binding protein (CRH-­BP) also exists, and for most of pregnancy it is present in excess of CRH in the maternal circulation. As CRH-­BP binds CRH with greater affinity than the CRH receptor, it is thought to sequester and therefore suppress CRH activity. Thus, for most of the pregnancy, the bulk of the placental CRH is considered to be inactive. However, during the last 4 weeks of pregnancy CRH-­BP levels decrease markedly.106 This coincides with the exponential increase in placental CRH production, which could result in a dramatic increase in CRH bioavailability (Fig. 11.4).

11

260

−90

−60

−30

0

Time from delivery (days) Fig. 11.4 Levels of corticotropin-­releasing hormone (CRH) and CRH binding protein (CRH-­BP) in the maternal circulation during human pregnancy. (From McLean M, Bisits A, Davies J, et al. A placental clock controlling the length of human pregnancy. Nature Med. 1995;1:460.)

Despite the elevated concentrations of CRH during pregnancy, secretion of ACTH from the maternal pituitary does not increase concordantly. In fact, pituitary ACTH levels remain low throughout pregnancy. The lack of CRH stimulation could be due to inhibition by the CRH-­BP. However, maternal ACTH production remains low late in gestation when CRH increases and CRH-­BP decreases. In vivo studies have shown that responsiveness of the maternal pituitary to CRH is markedly attenuated during pregnancy, and in vitro studies have shown that CRH downregulates CRH-­R expression in pituitary corticotropes.107 In vitro studies indicate that agents that increase CRH production by the hypothalamus also increase CRH production by placental cells.108 These agents include prostaglandins (PGs) E2 and F2α (PGE2 and PGF2α), norepinephrine, acetylcholine, vasopressin, angiotensin-­II, oxytocin (OT), interleukin (IL)-­1, and neuropeptide-­Y. In contrast, progesterone and nitric oxide donors inhibit placental CRH expression in vitro.108,109 Production of CRH by the placenta is increased by cortisol. This is in contrast to hypothalamic CRH, which is decreased by cortisol via a classic negative feedback loop. This stimulatory action has been observed in vivo in women who receive glucocorticoid treatment during the third trimester,110,111 and in vitro in cultured cytotrophoblast cells.112 The stimulation of placental CRH production by cortisol may result in a positive feedback endocrine loop. Placental CRH may stimulate ACTH production by the fetal pituitary, which would increase cortisol secretion by the fetal adrenals. Fetal adrenal cortisol could then further stimulate placental CRH production. The marked rise in placental CRH during the last 10 weeks of pregnancy could be due to such a positive feedback interaction and this endocrine loop may be involved in the process of parturition (discussed later in the chapter). CRH also influences fetal adrenal steroidogenesis by directly increasing dehydroepiandrosterone sulfate (DHEA-­S) production113,114 (Fig. 11.5). The capacity for CRH to act as an adrenal androgen secretagogue also has been demonstrated in vivo in adult men.115 Placental CRH and fetal adrenal DHEA-­S increase concordantly during the third trimester. The placental CRH-­ fetal adrenal endocrine axis may play a key role in the regulation of human parturition (discussed later in the chapter). Several actions have been ascribed to placental CRH in the control of human pregnancy. CRH may serve an autocrine-­ paracrine function within the placenta by regulating the expression and processing of POMC.116 Placental CRH may be part of the fetal-­placental stress response mechanism. The placenta is comparable to the hypothalamus in its production of CRH in response to stress. Neurotransmitters and neuropeptides activated in response to stress stimulate placental CRH release in vitro.117,118 The physiological implications of this are that the fetus may mount a stress response via placental CRH. This may be critical in conditions such as preeclampsia, placental vascular insufficiency, and intrauterine infection.

20 0 ACTH (1 nM)

−180 −150 −120

40

0.1

ACTH (1 nM)

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60

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1000

80

Control

2000

Cortisol (pmol/100K cells/24 h)

3000

0.1

1

10 100 CRH (pM)

1000 10,000

1

10 100 CRH (pM)

1000 10,000

500 DHAS (nmol/100K cells/24 h)

CRH or CRH-BP (pmol−1)

PART I  The Fundamentals of Reproduction

400 300 200 100 0

Fig. 11.5 Effect of corticotropin-­releasing hormone (CRH) on cortisol and DHEA-­S production in cultured human fetal (midgestation) adrenal cortical cells. (Adapted from Smith R, Mesiano S, Chan EC, Brown S, Jaffe RB. Corticotropin-­releasing hormone directly and preferentially stimulates dehydroepiandrosterone sulfate secretion by human fetal adrenal cortical cells. J Clin Endocrinol Metab. 1998;83:2916–2920.)

The CRH family of neuropeptides includes a group of structurally and functionally related proteins known as urocortins119,120 that have a high affinity for both CRH receptors. The urocortins are expressed by the syncytiotrophoblast and extravillous cytotrophoblasts and by the decidua and fetal membranes.121,122 Circulating levels of urocortins are low compared with CRH during pregnancy.123 Urocortin stimulates ACTH and PG production by trophoblast cells and causes vasodilation of the uteroplacental vasculature via activation CRH-­R2.124–126 In women with decreased uterine artery blood flow during midgestation, circulating urocortin levels are reduced in proportion to the increase in uterine artery resistance.127 In vitro studies suggest that this activity is compromised in preeclampsia.128 Studies in animal models also show vasodilatory effects of urocortin on placental blood flow and suggest that one of its main functions is to protect the fetus from hypoxic insults.129,130 Studies of term myometrial cells show that urocortin acting via CRH-­R2 increases contractility suggesting that it plays a role in parturition.131

POMC Derivatives The human placenta expresses POMC.132 In pituitary corticotropes, this 31-­kDa glycoprotein is the precursor for the ACTH-­ endorphin family of peptides. POMC is enzymatically cleaved to produce ACTH, β-­lipotrophic hormone (β-­LPH), α-­melanocyte-­ stimulating hormone (α-­MSH), and β-­endorphin (β-­EP). These neuroendocrine hormones play major roles in the physiological response to stress and the control of behavior. Each of these peptides, including full-length POMC, has been detected in the

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

The syncytiotrophoblast expresses POMC in a transcriptional pattern similar to that of extrapituitary tumors.135 However, the processing of POMC in the placenta is different than that in the pituitary. Although some placental POMC is cleaved, a significant amount of intact POMC is secreted by the placenta into the maternal circulation. In contrast, POMC is processed completely in the pituitary and in nonpregnant adults is undetectable in the circulation.134 However, during pregnancy, maternal circulating POMC levels are readily detectable by the third month and then increase steadily until midgestation, reaching a plateau of around 300 U/mL between 28 weeks and term. Soon after birth, POMC returns to undetectable levels. Unlike its pattern of secretion by the anterior pituitary, POMC produced by the placenta has no diurnal variability and it is not inhibited by glucocorticoids. Interestingly, during the third trimester maternal POMC levels closely correlate with plasma CRH levels but do not correlate with plasma ACTH or cortisol levels.134 The physiological role, if any, of placental ACTH and other POMC-­ derived proteins in the control of human pregnancy remains to be elucidated. With regard to fetal adrenal growth, placental ACTH plays a negligible role, because it is not sufficient to prevent adrenal hypoplasia in fetal hypopituitarism due to anencephaly. However, placental ACTH may influence maternal physiology and could be responsible for the relative resistance to negative feedback suppression of pituitary ACTH by glucocorticoids during pregnancy.136 Other POMC products are produced by the human placenta. Immunoreactive β-­EP in the maternal circulation remains relatively low throughout pregnancy and rises during labor and delivery indicative of the stress of parturition. Factors that increase pituitary ACTH (e.g., hypoxia and acidosis) also increase β-­EP production.137,138 Endogenous opioids, enkephalins, and dynorphins are also produced by the placenta. Immunoreactive methionine-­ enkephalin has been found in the human placenta. Circulating levels of methionine-­ enkephalin do not change appreciably throughout pregnancy.139 Three forms of dynorphin have been found in the human placenta.140 The amount of dynorphin in the placenta at term is similar to that found in the pituitary gland and brain. Relatively high concentrations of dynorphin are detectable in amniotic fluid and umbilical venous plasma, and maternal plasma levels in the third trimester, and at delivery are higher than in nonpregnant women.140 Dynorphin binds to kappa opiate receptors, which are abundant in the human placenta. Dynorphin receptor agonists stimulate the release of placental lactogen (PL; see below)141 suggesting that dynorphin exerts local regulatory effects on PL production.

Thyrotropin-­Releasing Hormone A substance similar to the hypothalamic thyrotropin-­releasing hormone (TRH) has been found in the human placenta.85 It stimulates the release of pituitary thyroid stimulating hormone (TSH) release in the rat both in vitro and in vivo, but is not identical to hypothalamic TRH.142 To date, a placental TSH has not been identified. Whether placental TRH plays a role in stimulating fetal or maternal pituitary TSH remains to be ascertained. The thyroid-­stimulating activity of the placenta has been ascribed to CG.

Placental Somatotropins In most eutherian mammals the placenta expresses members of the growth hormone (GH)-­PL gene family.143 The genes are encoded by a 66 kb segment of chromosome 17 that includes five closely related genes: GH1, GH2, CSH1, CSH2, and CSHL1, each derived from the duplication of a common ancestral gene. GH1 encodes pituitary GH and is expressed only in the pituitary. The

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Fig. 11.6 Levels of PL in the maternal circulation in relation to placental weight (Pl wt). (From Selenkow HA, Saxena BM, Dana CL, Emerson K Jr. Measurement and pathologic significance of human placental lactogen. In: Pecile A, Fenzi C, eds. The Foeto-­Placental Unit. Ecerpta Medica. Elsevier Science Publishers; 1969:340–362.)

other four are expressed exclusively in the placenta. GH2 encodes a placental GH (PGH) variant, which differs from pituitary GH by 13 amino acids. CSH1 and CSH2 are identical and encode PL. Placental lactogen Placental lactogen is a single-­chain polypeptide of 191 amino acids with 96% homology with GH. It can be detected in the placenta from around day 18 of pregnancy and in the maternal circulation by the third week of pregnancy. Low levels of PL (7–10 ng/mL) are present in the maternal circulation by 20 to 40 days of gestation. Thereafter, PL levels in the maternal circulation increase exponentially, reaching levels of 5 to 10 μg/mL at term.144 In contrast to the elevated levels of PL in the maternal circulation, concentrations of PL in the fetal circulation range from 4 to 500 ng/mL at midgestation and only 20 to 30 ng/mL at term.145 Thus, PL is preferentially secreted into the maternal compartment. In normal pregnancy, PL is first synthesized by the cytotrophoblasts of the developing placenta during the first 6 weeks of pregnancy. Thereafter, expression switches to the syncytiotrophoblast, which eventually becomes the exclusive source of PL. The extent of PL expression by the syncytiotrophoblast does not change during the course of pregnancy, although total placental production increases substantially.146 Therefore, the rise in PL production is thought to be due to the increase in placental mass. Maternal PL levels rise concordantly with an increased amount of syncytiotrophoblast tissue as gestation advances (Fig. 11.6). After delivery of the placenta, the half-­life of the disappearance of circulating PL is 9 to 15 minutes. To maintain circulating concentrations, this would imply placental production of 1 to 4 g of the hormone per day at term.147 Thus, the production of PL represents one of the major metabolic and biosynthetic activities of the syncytiotrophoblast. PL is expressed by all types of trophoblastic tissue and has even been detected in the urine of patients harboring trophoblastic tumors, in men with choriocarcinoma of the testis, and in the serum and urine of women with molar pregnancies. Factors that regulate PL production have been assessed in cultured cytotrophoblast cells. Insulin and growth hormone-­ releasing factor (GHRF) stimulate PL secretion, whereas somatostatin (SS) inhibits its secretion. The presence of PL, PGH, SS, and GHRF in the same cell suggests that another autoregulatory loop analogous to the hypothalamic-­pituitary axis operates within the placenta. In the third trimester, maternal PL and GHRF Interestingly, SS expression is

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maximal in early pregnancy and decreases during the second and third trimesters, a pattern opposite to that of PL.149 Thus, locally produced GHRF and SS may regulate placental PL production. Several studies have demonstrated changes in maternal PL levels in response to metabolic stress. Specifically, prolonged fasting at midgestation150 and insulin-­induced hypoglycemia raise maternal PL concentrations.151 However, PL levels do not change in association with normal metabolic fluctuations during a typical 24-­hour period.152 The initial identification of PL was based on its lactogenic activity in bioassays, suggesting that PL acts as a lactogen in human pregnancy. Indeed, PL binds to the prolactin receptor with relatively high affinity (Kd 0.1 nM). In contrast, its affinity for the GH receptor is lower (Kd 770 nM).153 Thus, PL may function mainly as a lactogen during pregnancy with only minimal activity as a somatogen (promotes growth). However, administration of PL to nonpregnant women in sufficient quantities to mimic pregnancy levels did not induce lactation.154 This does not rule out PL as a lactogen, as its actions on the mammary gland may be dependent on other factors in the endocrine milieu of pregnancy (e.g., estrogens and progesterone). Nonetheless, the in vivo lactogenic properties, if any, of PL in human pregnancy remain to be established. It should be noted that maternal prolactin levels increase significantly in the later stages of pregnancy and, together with estrogen and progesterone, are likely sufficient to induce mammary growth and lactation. Placental Growth Hormone. Two forms of PGH have been identified in the human placenta, both of which are expressed by the syncytiotrophoblast.155 The smaller, 22 kDa form is almost identical to pituitary GH, differing by only 13 amino acids. The larger 26kDa PGH is a splice variant that retains intron 4. The extent of PGH production is significantly less than that of PL, and PGH is not secreted into the fetal compartment.156 Levels of PGH in the maternal circulation are approximately 1000-­ fold less than PL and can be detected only after midgestation (between 21 and 26 weeks). During the third trimester, maternal PGH levels increase exponentially, in concert with PL, and reach a maximum of approximately 20 ng/mL by term. In the first trimester, pituitary GH is measurable and secreted in a highly pulsatile manner.157 However, pituitary GH production decreases progressively from about week 15, and by 30 weeks cannot be detected. During the same period, nonpulsatile secretion of PGH by the placenta increases markedly and becomes the dominant GH of pregnancy. Function of PL and PGH. Studies of PL and PGH deficiency have revealed their potential synergistic roles in human pregnancy.158 In all cases of complete PL deficiency (i.e., no detectable PL in maternal blood or in the placenta) pregnancy and fetal development were normal. However, deficiency of both PL and PGH due to a mutation in the GH/PL gene cluster is associated with severe fetal growth retardation. Pregnancies in which only PGH is deficient have not been identified. These experiments of nature indicate that PL is not necessary for normal pregnancy, whereas PGH is an important regulator of fetal growth. As PGH does not enter the fetal compartment, it likely influences fetal growth via effects on the mother. This is consistent with the thesis that PL and PGH modulate maternal metabolism to meet fetal energy requirements.159 Maternal food intake and intestinal calcium absorption increase during the first trimester, and insulin secretion immediately after feeding almost doubles. In the second half of pregnancy, maternal cells become increasingly resistant to insulin, i.e., a diabetogenic state, that is thought to be promoted by the combined GH-­like and contra-­insulin activity of PGH and PL. This decreases glucose uptake and increases free fatty acid release. The net effect is increased availability of free fatty acids, glucose,

and amino acids for fetal consumption. Decreased glucose mobilization into maternal cells would increase the supply of glucose available for the fetus, which is especially important for fetal brain growth and development. Free fatty acids can cross the placenta, and the increased ketones induced by their metabolism are also an important energy source for the fetus. Thus, during the second half of pregnancy PGH and PL direct maternal metabolism toward the mobilization of maternal energy resources to furnish the needs of the developing fetus.160 Soon after birth, insulin resistance reverts to the normal nonpregnant state, suggesting that maternal glucose homeostasis is influenced by hormonal factors produced by the fetus-­placenta. This fetal-­maternal hormonal interaction represents an example of genetic conflict.161 The fetus, through natural selection, acquires traits, for example, PGH and PL, that favor the extraction of resources from the maternal compartment. Conversely, mothers have evolved mechanisms to counteract fetal demand. Disorders on either side of this equation lead to pathophysiology. For example, in women with gestational diabetes, insulin secretion is insufficient to balance the decrease in insulin sensitivity and consequently blood glucose levels increase leading to an oversupply of glucose in the fetus resulting in macrosomia.

Growth Factors The human placenta produces many growth factors and cytokines.70,162–164 Placental growth factors are thought to be involved in the control of implantation, angiogenesis, and vascularization of the implantation site, and the establishment and growth of the placenta. Some important growth factors produced by the human placenta are discussed below. Insulin-­like growth factors. The human placenta produces IGF-­ I, expressed by the syncytiotrophoblast, and IGF-­ II, expressed by cytotrophoblasts, the syncytiotrophoblast, and extravillous trophoblasts, from as early as week 8 of human pregnancy.165,166 Gene deletion studies in mice have shown that IGF-­I and IGF-­II are key regulators of placental and fetal growth. Pups of IGF-­II knockout mice exhibit fetal growth restriction but normal growth after birth, whereas pups of IGF-­I knockout mice have poor postnatal growth and die before reaching adulthood.167 Specific inhibition of placental IGF-­II expression inhibits placental and fetal growth168,169 demonstrating the key role of placental IGF-­II during fetal development. IGF-­II is an imprinted gene that is expressed only by the paternal allele.170 In general, genes expressed only by the paternal allele promote fetal/placental growth, whereas those expressed only by the maternal allele suppress fetal growth.171 Loss of IGF-­ II imprinting is associated with fetal growth restriction that is thought to be secondary to placental dysfunction.172 In Beckwith-­Wiedemann syndrome, abnormal methylation of the imprinting centers on chromosome 11 leads to expression of IGF-­II from the maternal allele, which is normally suppressed. The extra IGF-­II produced in this condition causes fetal macrosomia.173 The finding that a paternal gene promotes placental growth further reflects the concept of genetic conflict between the maternal and fetal genomes.161 The size and ultimate health of the fetus depend greatly on the size of the placenta. Growth factors that increase placenta size are an advantage to the fetus because they allow it to extract resources more efficiently from the mother. Passage of paternal genes to the next generation is favored if the nutrient supply to the fetus is maximized. Maternal genes, on the other hand, not only must survive to the next generation, but they also must ensure that the current pregnancy does not compromise the mother’s future reproductive capacity. Maternal genes would therefore be selected to oppose the effects

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

IGFs in the circulation are bound to six binding proteins (IGF-­BPs) that control IGF function by sequestering circulating IGFs and controlling their bioavailability to interact with receptors on target cells. During pregnancy, IGFBP-­1 is produced by the decidua and all of the IGFBP-­1 in amniotic fluid is maternally derived.169 Overexpression of decidual IGFBP-­1 inhibits placental and fetal growth primarily by sequestering placental IGF-­II.169 Epidermal growth factor family. The epidermal growth factor (EGF) family includes EGF, heparin-­binding EGF-­like growth factor (HB-­EGF), amphiregulin, betacellulin, epiregulin, and TGFα. HB-­EGF is expressed by the syncytiotrophoblast and cytotrophoblast cells and this is relatively high in the first trimester and decreases with advancing gestation.174 EGF-­family growth factors are thought to promote trophoblast invasion during implantation and deficiency in EGF expression and/ or signaling is associated with preeclampsia and fetal growth restriction.175,176 Vascular endothelial growth factor family. The vascular endothelial growth factor (VEGF) family of peptides comprises VEGF-­A (referred to as VEGF), VEGF-­B, VEGF-­C, VEGF-­D, VEGF-­E (generated by alternative splicing from a single gene), and 4 splice variants of a factor produced by trophoblast cells known as placenta growth factor (PlGF).177–181 These peptides are mitogenic and antiapoptotic for endothelial cells and promote the formation of new blood vessels via angiogenesis and vasculogenesis, and increase tissue perfusion via vasodilatation and increased microvascular permeability.182,183 The actions of VEGFs and PlGFs are mediated by two tyrosine kinase receptors FMS-­like tyrosine kinase-­1 (FLT-­1) and kinase insert domain receptor (KDR).184–186 A soluble form of FLT-­1 (sFLT-­1) also exists, which is present in the circulation and binds VEGFs and PlGFs to neutralize their angiogenic activity.187 Remodeling of the uterine spiral arterioles, and continued angiogenesis through pregnancy to match placental growth with appropriate maternal vascular supply, are critical for the success of pregnancy. These events are affected by VEGFs and PlGFs expressed by cytotrophoblasts, the syncytiotrophoblast and villous stromal cells in the placenta; and VEGF/PlGF receptor expressed by endothelial cells at the placental/maternal interface.188–190 The placental VEGF/PlGF system is involved in pregnancy complications, especially preeclampsia, which is caused by placental factors affecting maternal hemodynamic homeostasis.191 The etiology of preeclampsia is thought to involve placental hypoxia, due to decreased invasion of cytotrophoblasts into the maternal spiral arteries early in pregnancy that compromises placental perfusion later in gestation.192,193 This is thought to induce compensatory mechanisms involving the VEGF/PlGF system to increase placental perfusion by modulating maternal hemodynamics. One mechanism for this is that hypoxia increases placental expression of VEGF which, via KDR, increases trophoblast expression of sFLT1194 leading to increased levels of sFLT1 in the maternal circulation. In pregnancies complicated with preeclampsia, circulating sFLT1 levels in the mother are increased compared with normal pregnancies.195–197 Consequently, sFLT1 levels have biomarker potential to assess risk for preeclampsia.198,199 Excess sFLT1 in the maternal circulation may disrupt maternal endothelium function leading to the symptoms of preeclampsia, especially hypertension and proteinuria.200 Thus, the placental VEGF/PlGF system appears to play a key role in maintaining maternal vascular function to provide sufficient placental perfusion necessary for normal fetal/ placental development. VEGF/PlGF-­ mediated compensatory mechanisms in response to inadequate placental perfusion may adversely affect the maternal vasculature leading to complica tions such as preeclampsia.

263

Fibroblast growth factor family. Like VEGF, the fibroblast growth factor (FGF) family (comprises 23 members: FGF-­1 to -­ 23) of polypeptides are mitogens that affect endothelial cell migration and promote the formation of blood vessels.201 Effects of FGFs are mediated by four specific high-­affinity cell surface FGF receptors (FGFR-­1 to -­4). The human placenta expresses mainly FGF-­2 and FGF-­10 that localizes to villous trophoblasts and connective tissue stroma of mesenchymal villi.201,202 Interestingly, each of the FGFRs are expressed by Hofbauer cells,201,203 placental villous macrophages of fetal origin that are thought to play an important role in placental development including vasculogenesis and angiogenesis. FGF induces Hofbauer cells to produce multiple growth factors and cytokines involved in tissue repair. Activation of Hofbauer cells by FGF may promote placental growth by facilitating the outgrowth of syncytiotrophoblast buds.204,205 Adipokines. Adipokines are factors produced by adipose tissue that affect metabolic homeostasis, satiety, and reproduction. Currently, known adipokines include leptin, adiponectin, resistin, ghrelin, and visfatin. Adipokines produced by the placenta regulate the maternal metabolic adaptation to pregnancy, especially increased insulin resistance.206 Leptin, a 146-­amino acid protein produced primarily by adipocytes, is a key regulator of satiety and body mass index and its levels are thought to reflect the amount of energy stores and nutritional state.207 Leptin decreases food intake and body weight via its hypothalamic receptor.208 In the reproductive system, leptin is thought to coordinate body mass status with reproductive function.209 In general, leptin acts as a permissive factor. Pulsatile hypothalamic GnRH secretion does not occur unless leptin levels reach a threshold value. Such a mechanism may ensure that energy stores are sufficient to support a pregnancy. The placenta is the principal source of leptin during pregnancy. Leptin produced by the placenta is secreted mainly into the maternal circulation, and as a consequence, leptin levels are elevated during pregnancy. In the first trimester, maternal plasma leptin levels are double nonpregnant values and continue to increase during the second and third trimesters,210 during which time it is also expressed by the chorion and amnion.211 Leptin levels decline to normal nonpregnant values within 24 hours of delivery.212,213 The influence of placental leptin on maternal biology is unclear. Abnormally high placental leptin production is associated with maternal diabetes mellitus and hypertension, and umbilical leptin levels correlate with fetal adiposity.214 Interestingly, leptin levels during pregnancy do not correlate with body mass index as they do in the nonpregnant state. Pregnancy appears to be a state of hyperleptinemia and leptin resistance, with uncoupling of leptin effects on eating behavior, satiety, and metabolic activity.215 Leptin is lipolytic and favors fatty acid mobilization from adipose tissue. It also may act in the liver, pancreas, and muscle to decrease insulin sensitivity and mobilize glucose. The human placenta expresses leptin receptors and therefore leptin can act in a paracrine manner to modulate placental function. Leptin induces CG production in trophoblast cells and is also thought to increase placental growth by augmenting mitogenesis, amino-­acid uptake, and extracellular matrix synthesis.216,217 Adiponectin, resistin, ghrelin, and visfatin are also produced by the placenta and secreted into the maternal and fetal compartments. As with leptin, these adipokines appear to affect maternal metabolic homeostasis in favor of pregnancy and the supply of nutrients to the fetus.213,218–220

Steroid Hormones and the Fetal-­Placental Unit The human placenta is a steroidogenic organ. For most of the pregnancy, it produces progesterone and estrogens. Steroid hormone production by the placenta is dependent on precursors

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Acetate

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Fig. 11.7 Biosynthetic pathways in steroid hormone formation. The C-­21 compounds include progestins and adrenal corticosteroids. The C-­19 compounds include androgens, and the C-­18 compounds include estrogens. Steroids with a double bond between the 5 and 6 positions in the steroid nucleus (Δ5-­steroids) are shown on the left, and those with a double bond between the 4 and 5 positions (Δ4-­steroids) are depicted on the right. The principal steroid metabolizing enzymes are shown in italics.

provided by the fetus and mother. The interdependence of fetus, placenta, and mother for steroid hormone production in human pregnancy led to the concept of an integrated fetal-­placental-­ maternal steroidogenic unit. To understand this concept, it will be useful to review the general steroid biosynthetic pathways (Fig. 11.7) (also discussed in Chapter 4). Progesterone. The human placenta produces large amounts of progesterone throughout pregnancy. It does this mainly by converting low-­ density lipoprotein cholesterol, extracted from the maternal circulation, to pregnenolone, which is then converted to progesterone.170 Cholesterol is converted to pregnenolone in the mitochondria of cytotrophoblast cells and the syncytiotrophoblast by the cytochrome P450scc enzyme. This is the rate-­determining step for placental progesterone production. The conversion of pregnenolone to progesterone also occurs in the mitochondria and is catalyzed by type-­ 1 3β-­ hydroxysteroid dehydrogenase (3βHSD-­ I). The human placenta is highly inefficient at converting progesterone to 17α-­ hydroxyprogesterone.221 Although the human placenta expresses P450c17, albeit at low levels, and has an intact Δ4 steroidogenic pathway,222 the efficiency of progesterone synthesis is far greater than its metabolism through the Δ4 pathway, resulting in a net accumulation and secretion of progesterone. Production of progesterone approximates 250 mg/day by the end of pregnancy, at which time circulating levels are on the order of 130 ng/mL.223 As its name implies, progesterone is a progestation hormone. It is essential for the establishment and maintenance of pregnancy. In all viviparous animals examined so far, disruption of progesterone synthesis or action during pregnancy induces labor and delivery. The role of progesterone in the maintenance of pregnancy and the mechanism by which its actions are withdrawn to trigger parturition are discussed later in this chapter. Estrogens. The principal roles of estrogens in human pregnancy are to stimulate uterine growth and increase uterine blood flow. Estrogens also affect breast development in preparation for lactation. At parturition, estrogens oppose the actions of progesterone by augmenting uterine contractility and inducing cervical softening. Estrogens are synthesized by the human placenta from C19 steroids.224 The principal precursor used for placental

estrogen formation is dehydroepiandrosterone (DHEA) sulfate (DHEA-­ S), supplied mainly by the fetal adrenal glands. Because the placenta has an abundance of the sulfatase (sulfate-­ cleaving) enzyme, DHEA-­S is rapidly converted to free (unconjugated) DHEA, which is then converted to androstenedione by 3βHSD-­I. The human placenta also expresses high levels of the aromatase enzyme, which converts androstenedione to estrone. The 17β hydroxysteroid dehydrogenase (17βHSD) enzymes then interconvert estrone and estradiol (Fig. 11.8). The major estrogen formed during human pregnancy is estriol, which has an additional hydroxyl group at position 16. Estriol constitutes more than 90% of the estrogen in pregnancy urine, into which it is excreted as sulfate and glucuronide conjugates. Estriol production by the placenta increases with advancing gestation and ranges from approximately 2 mg/24 hours at 26 weeks to 35 to 45 mg/24 hours at term.225 At term, the concentration of estriol in the maternal circulation is 8 to 13 ng/dL.226 In contrast, ovarian production of estriol in nonpregnant women is barely detectable.226 Placental estriol is formed by a biosynthetic process unique to human (and some higher primates) pregnancy, that begins with DHEA-­S from the fetal adrenal glands. When DHEA-­S of either fetal or maternal origin reaches the placenta, estrone and estradiol are formed. However, little of either is converted to estriol by the placenta. Instead, some of the DHEA-­S undergoes 16α-­ hydroxylation to form 16α-­OH-­DHEA-­S, primarily in the fetal liver and, to a limited extent, in the fetal adrenal. In the placenta, that sulfatase enzyme converts 16α-­OH-­DHEA-­S to 6α-­OH-­ DHEA, which is then aromatized to form estriol. Thus, estriol in the maternal blood reflects the steroidogenic activity of the fetal hypothalamic-­pituitary-­adrenal (HPA) axis. In the mother, estriol is conjugated to form estriol sulfate and estriol glucosiduronate in the liver and excreted via the urine.227 The function of estriol in pregnancy has attracted much speculation. In most biological systems, estriol is a weak estrogen with approximately 1% the potency of estradiol and 10% that of estrone. However, in pregnancy, uterus estriol is as effective as the other estrogens in increasing uteroplacental blood flow.228 Another placental estrogen derived from a fetal precursor is estetrol, formed after 15-­hydroxylation of DHEA-­S. Its function is not known. Relative levels of progesterone and estrogens (estrone, estradiol, estriol, and estetrol) in the maternal circulation during human pregnancy

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

Mother

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Fig. 11.8 Biosynthesis of progesterone and estrogens by the human placenta. Progesterone is produced mainly from maternal cholesterol. P450c17 is not expressed in the human placenta and therefore, progesterone cannot be converted to C19 androgens. Instead, estrogens are biosynthesized from C19-­androgen precursors (mainly DHEA-­S) provided by the maternal and fetal adrenals. (From Mesiano S. Roles of estrogen and progesterone in human parturition. Front Horm Res. 2001;27:86.)

Production of estrogens by the human placenta is extraordinarily high and circulating levels during pregnancy are several orders of magnitude higher than physiological levels in nonpregnant women. A key question, therefore, is why does the human placenta make so much estrogen? Conditions in which placental estrogen synthesis is markedly decreased (e.g., anencephaly, congenital adrenal lipoid hyperplasia, placental aromatase deficiency, and placenta sulfatase deficiency) have provided some insight into the role of the placental estrogens in human pregnancy. Although pregnancy was prolonged in some cases of placental sulfatase deficiency229–232 and anencephaly,233–236 in most cases a marked decrease in placental estrogen synthesis had little effect on fetal and placental development and the timing of parturition. Likewise, pregnancies with lowered estrogen levels due to placental aromatase deficiency,237,238 appear normal, although mothers and female fetuses exhibited virilization due to excessive amount of DHEA that has weak androgenic activity. Thus, high levels of estrogens of placental origin may not be essential for normal pregnancy and parturition, but rather a critical role of placental aromatase is to metabolize circulating androgens to protect the female fetus and mother from virilization. These observations do not exclude the role of estrogen in the control of human pregnancy. Although levels of maternal estrogens in pregnancies with placental estrogen synthesis defects were low compared with normal pregnancies, they were still in a physiologically significant range (1–1.6 nmol/L)237 238 and com parable to levels reached in the midcycle and luteal phase of the

4

8 12 16 20 24 28 32 36 40 Gestational Age (weeks)

Fig. 11.9 Schematic depiction of maternal progesterone and estrogen (estradiol, estrone, and estriol) concentrations through human pregnancy compared with average levels in normal cycling women.

menstrual cycle (0.6–2 nmol/L). Thus, despite the inability of the placenta to produce estrogens in certain abnormalities such as aromatase deficiency, the estrogen target tissues were still exposed to moderately high levels of estradiol. This suggests that a minimal level of estrogen is necessary for human pregnancy and that the excessive levels produced by the placenta are redundant. Studies in the baboon placenta suggest that estrogen upregulates 11β-­HSD-­2 expression in the placenta. Because estrogen synthesis by the placenta requires fetal adrenal androgen precursors (produced in part by fetal pituitary ACTH stimulation), the estrogen regulation of placental 11β-­HSD-­2 represents a regulatory loop that ensures that maternal cortisol does not affect the fetal HPA axis.239 For most of gestation, the placenta expresses the 11β-­ hydroxysteroid dehydrogenase type 2 (11β-­HSD-­2) enzyme, which inactivates cortisol by catalyzing its conversion to cortisone.240,241 As maternal cortisol levels are three times those of the fetus, this biochemical barrier serves to prevent excess cortisol from entering the fetal compartment. This is important because exposure of the fetus to high levels of cortisol not only could interfere with the fetal HPA axis but also is associated with decreased birth weight and hypertension.242–244

Placental Extracellular Vesicles Villous trophoblasts and the syncytiotrophoblast produce extracellular vesicles (EVs; exosomes) that contain biologically active signaling factors in the form of nucleic acids (messenger RNA, micro RNA, and noncoding RNA), proteins, and lipid;245 and as such can be considered vehicles to transport hormones and signaling molecules between cells. Exosomes (30–100 nm vesicles) are produced by trophoblast cells and shed via endosomal trafficking primarily into the maternal compartment.245,246 Exosomes

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are thought to exert biological effects by fusing with the plasma membrane of target cells and depositing their cargo of bioactive factors into the cytoplasm.247 Placental exosomes may regulate proximal and distal targets and could be key vectors for communication from the fetus/placenta to the mother.248,249 They can be detected in the maternal circulation from the 6th week of gestation and their levels increase gradually with advancing gestation in proportion to the increased size of the placenta.245 Levels of exosomes (placental and nonplacental) in the maternal circulation and the composition of their cargo have been associated with pregnancy complications such as preeclampsia, preterm birth, intrauterine growth restriction, and gestational diabetes.246,250 This suggests that the content of placental exosomes could be used as biomarkers to predict specific pregnancy complications.251 Placental exosomes also could exert physiologic effects. mi-RNAs in placental exosomes affect the function of local immune cells to boost resistance to viral infection252 In addition, placental exosomes carry factors to the maternal compartment that affect maternal immune tolerance toward the fetus at the maternal-­fetal interface.253–255

Cell-­Free Fetal DNA Trophoblast EVs also contain fragmented (∼200 base pairs in length) fetal DNA, referred to as cell-­free fetal DNA (cffDNA). cffDNA originates in trophoblast cells and is detectable in the maternal circulation from around the 7th week of gestation. cffDNA increases with advancing gestation such that late in pregnancy it represents ∼4% of the cell-­free DNA.256–258 After delivery of the placenta, the amount of cffDNA in maternal blood rapidly decreases. The physiologic role of cffDNA and why it is packaged into vesicles that are shed from trophoblast cells into the maternal compartment is not known. Because it can be isolated and sequenced, cffDNA is used for noninvasive prenatal diagnosis and testing for conditions such as X-­linked genetic disorders and aneuploidies as well as fetal sex determination.259

FETAL NEUROENDOCRINE SYSTEMS • Appropriate development of the fetal neuroendocrine system is critical for its survival as a neonate. • Functional maturation of neuroendocrine systems is necessary for the newborn to establish and maintain homeostasis. • The future health of the newborn may be influenced by the development trajectory of the neuroendocrine system during fetal life.    The appropriate functional maturation of neuroendocrine systems during fetal life has profound effects on long-­term health into adulthood.260 This phenomenon, referred to as developmental origins of adult health and disease, was first observed in epidemiological studies showing an association between perturbations of intrauterine nutritional status and the development of hypertension, insulin resistance, and obesity that predispose to cardiovascular disease, diabetes, and the metabolic syndrome in adulthood.261 The developmental origins hypothesis posits functional plasticity in the development of fetal neuroendocrine and organ systems such that a single genotype gives rise to multiple “normal” phenotypes in response to environmental cues (e.g., extent of maternal nutrition and stress). Thus, the fetus prepares itself for the extrauterine environment by modulating its physiological systems to match the anticipated extrauterine environment based on maternal cues. Interestingly, neuroendocrine plasticity does not continue later in life. The period of neuroendocrine plasticity is limited to fetal and possible early neonatal development and the physiological changes appear to be permanent. For example, studies in animals show that changes in basal and

stress-­associated fetal HPA activity induced in utero by maternal starvation persist after birth. The altered stress-­response pathway may underpin pathophysiological conditions that develop during adulthood.262,263 The physiological, biochemical, and genetic mechanisms that mediate the reactive plasticity of fetal neuroendocrine development are not clearly defined. The placenta is thought to play an active role, although it remains uncertain how information regarding the environment is transmitted from the mother, through the placenta, and to the fetus.264 As described above, the placenta produces a plethora of hormones that could influence the developmental trajectories of fetal neuroendocrine axes. For example, placental CRH may modulate the responsiveness of the fetal adrenal cortex to ACTH and therefore contribute to the establishment of the HPA set-­point.265 It is also possible that perturbations in the gestational milieu induce static changes in chromatin structure, referred to as epigenetic modifications, that alter gene expression and persist into adult life.266 The most common epigenetic modifications (also known as marks) are methylation of cytosines in CpG dinucleotides and posttranslational modifications (e.g., acetylation) of histones. Methylation in gene promoter regions generally inhibits downstream gene expression, and histone modifications alter the chromatin secondary structure, affecting the activity of the transcriptional machinery.267 These chemical modifications do not involve changes in the primary DNA sequence and are heritable by mitosis from one cell generation to the next.268 During the preimplantation period, epigenetic information inherited from the gametes is reset through waves of DNA demethylation in embryonic cells. Although most epigenetic marks are erased, methylation of imprinted genes is mostly preserved (see Chapter 9). As the embryo develops, the DNA is gradually remethylated. This provides an opportunity for environmental factors to influence the epigenome (genome-­wide pattern of epigenetic marks).269 Epigenetic modification during development is proposed as a mechanism for environment-­induced phenotypic plasticity and provides a mechanistic explanation for the effects of prenatal and perinatal environmental exposures (e.g., malnutrition) on the function of neuroendocrine systems and their effects on subsequent disease risk in later life.270–273 The current understanding of the developmental patterns of the major neuroendocrine systems in the human fetus is discussed below.

Hypothalamic Hormones By the end of the fifth week of pregnancy, the primitive hypothalamus can be identified as a swelling on the inner surface of the diencephalic neural canal. It then differentiates to form a complex of interconnecting nuclei. By 9 to 10 weeks, the median eminence of the hypothalamus is evident, and interconnecting fiber tracts of hypothalamic nuclei can be identified by 15 to 18 weeks. The hypophysiotropic hormones GnRH, TRH, CRH, GHRF, and SS appear in the fetal hypothalamus during this period (Table 11.2).

Gonadotropin-­Releasing Hormone The human fetal hypothalamus produces immunoreactive and bioactive GnRH by 10 weeks of gestation. Between 10 and 22 weeks, the concentration of GnRH in the fetal hypothalamus remains constant (0.27–13.1 pg/mg) and is not different between the sexes. GnRH release from human fetal hypothalamic explants is pulsatile.274,275 In male rhesus monkey fetuses, the pituitary-­ gonadal axis is active during the last third of gestation. The pituitary produces LH in response to GnRH and the testes produce testosterone in response to LH.276 A similar situation likely exists in the human male fetus. Whether this also is the case for the

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

TABLE 11.2  Ontogeny of Human Fetal Hypothalamic and Pituitary Hormones Hormone Hypothalamic Gonadotropin-­releasing hormone Thyrotropin-­releasing hormone Somatostatin Dopamine Growth hormone-­releasing hormone Corticotropin-­releasing hormone Pituitary Prolactin Growth hormone Corticotropin (ACTH) Thyroid-­stimulating hormone (thyrotropin) Luteinizing hormone Follicle-­stimulating hormone

Age Detected (wk) 14 10 14 11 18 16 16.5 10.5 7 13 10.5 10.5

Thyrotropin-­Releasing Hormone Significant levels of immunoreactive TRH are found in the human fetal hypothalamus early in gestation.274 As with GnRH, fetal hypothalamic TRH levels do not correlate with sex or gestational age. The presence of TRH in the fetal hypothalamus in early gestation and midgestation suggests its role in the regulation of TSH, and possibly PRL, secretion.

Growth Hormone-­Releasing Factor and Somatostatin GHRF can be detected in fetal hypothalamic neurons and fiber tracts at 18 weeks, with increasing levels found up to 30 weeks.277 The simultaneous detection of GHRF in neuron cell bodies and fiber tracts indicates its release into portal vessels by midgestation. Immunoreactive SS can be identified in the hypothalamus of human fetuses from 10 to 22 weeks.274 In contrast to GnRH and TRH, fetal hypothalamic SS increases with advancing gestation.

Corticotropin-­Releasing Hormone and Arginine Vasopressin CRH is a potent secretagogue for ACTH and β-­endorphin release by the human fetal pituitary gland.278 Arginine vasopressin (AVP) also directly stimulates ACTH secretion by the fetal pituitary and can synergize with CRH275. CRH-­immunoreactive fibers can be detected in the median eminence between 14 and 16 weeks.277,279 CRH and AVP have been detected in human hypothalamic extracts from 12 to 13 weeks.279 Hypothalamic CRH and AVP increase with gestational age, and the CRH bioactivity of fetal hypothalamic extracts, measured in isolated rat anterior pituitary cells, is augmented by AVP. Thus, the human fetal hypothalamus has the capacity to regulate pituitary ACTH production from early in the second trimester. The extent to which fetal hypothalamic CRH and AVP and placental CRH interact in regulating fetal ACTH release remains uncertain.

Catecholamine-­Dopamine Catecholamine in cells projecting from the arcuate nuclei to the internal and external layers of the median eminence appears during the interval from 12 to 16 weeks.280 Dopamine is present in the fetal hypothalamus at weeks 11 to 15 at a concentration twice that of the adult. Hypothalamic dopamine inhibits PRL release from the fetal pituitary during this time.281

Pituitary Hormones The pituitary primordium appears as the epithelial evagination of Rathke’s pouch arising from a diverticulum of the stomadeum.

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The evaginated pituitary primordium appears at 4 weeks and separates from the stomadeum by 5 weeks. The floor of the sella turcica is in place by week 7 and separates the pituitary from its epithelial origins. The progenitors of pituitary hormone-­ secreting cells originate in the ventral neural ridges of the primitive neural tube. This region also gives rise to the diencephalon, suggesting that the hypothalamus and anterior pituitary share a common embryonic origin. The pituitary increases in size and cell number through the proliferation of cell cords into mesenchyme beginning during week 6. Capillaries interdigitate among the mesenchymal tissue of Rathke’s pouch and the diencephalon at 8 weeks and the median eminence is distinguishable by 9 weeks. The hypothalamic-­hypophyseal vascular system in fetuses is intact by 11 to 16 weeks. The anterior pituitary gland comprises five types of specialized epithelium-­derived secretory cells: (1) lactotropes producing prolactin; (2) somatotropes producing GH; (3) corticotropes producing ACTH; (4) thyrotropes producing TSH; and (5) gonadotropes producing LH and FSH. Within the pituitary, cells containing ACTH have been detected at 7 weeks, β-­lipotropin and β-­endorphin at 8 weeks, GH-­ containing and LH-­ containing cells both at 10.5 weeks, and TSH-­containing cells at 13 weeks. Melanocyte-­stimulating hormone (MSH)-­ containing cells, which probably contain β-­ lipotropin, have also been reported to appear at 14 weeks, and prolactin-­containing cells appear at 16.5 weeks (Table 11.2).282 Many of the components required for the normal regulated function of the secretory cells of the anterior pituitary, the hypophysiotropic factors elaborated by the hypothalamus, and the neural-­vascular link connecting the hypothalamus and anterior pituitary, are present in the fetus well before the end of the first half of pregnancy. The primate fetal pituitary is competent to respond in vitro to virtually all of the known hypophysiotropic factors by midgestation. Whether it responds to these compounds in vivo remains to be demonstrated directly, although studies in the catheterized rhesus monkey fetus in utero suggest that responses to GnRH are intact at midgestation.276

Fetal Pituitary-­Adrenal Axis The anlage of the human adrenal cortex is first identified at about the fourth week of gestation as a thickening of the celomic epithelium in the notch between the primitive urogenital ridge and the dorsal mesentery. By the fifth week, these primitive cells begin to migrate toward the cranial end of the mesonephros, where they condense to form the earliest recognizable manifestation of the adrenal gland. Cells destined to become the steroidogenic cells of the adrenal and gonad are derived from neighboring areas of the coelomic epithelium and are morphologically identical. In general, the portion medial to the mesonephros produces cells destined for the adrenal cortex, whereas the portion ventral to the mesonephros produces cells destined for the gonad. During the last two-­thirds of gestation in humans and higher primates, the fetal adrenal glands are disproportionately enlarged and exhibit extraordinary growth and steroidogenic activity. The growth is attributable to cortical hypertrophy controlled by locally produced growth factors including basic FGF and IGF-­II in response to ACTH.283–285 For much of gestation, the human fetal adrenal cortex is composed of two morphologically distinct zones, the fetal zone and the definitive zone. The fetal zone accounts for 80% to 90% of the cortex and is the primary site of growth and steroidogenesis. During midgestation, the fetal zone produces 100 to 200 mg/day of DHEA-­S, which is quantitatively the principal steroid product of the human fetal adrenal gland throughout gestation. As its name implies, the fetal zone exists only during fetal life; it involutes soon after birth. Clusters of immature neuroblasts are

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present between the innermost fetal zone cells. These cells aggregate at birth to form a functional adrenal medulla. The definitive zone (also referred to as the adult cortex, neocortex, or permanent zone) comprises a narrow band of small, tightly packed basophilic cells that exhibit structural characteristics typical of cells in a proliferative state. Its inner layers form arched cords that send finger-­like columns of cells into the outer rim of the fetal zone. Definitive zone cells begin to resemble steroidogenically active cells later in pregnancy and appear to secrete mineralocorticoids late in the third trimester. Ultrastructural and functional studies have revealed a third zone between the fetal and definitive zones, referred to as the transitional zone, the cells of which have intermediate characteristics between the fetal and adult zones. After midgestation, transitional zone cells have the capacity to synthesize cortisol and thus may be analogous to cells of the zona fasciculata of the adult adrenal. By week 30 of gestation, the definitive zone and transitional zone begin to take on the appearance of the zona glomerulosa and the zona fasciculata, respectively. Thus, by late gestation, the fetal adrenal cortex resembles a rudimentary form of the adult adrenal cortex. The postnatal remodeling of the primate adrenal cortex involves a complex wave of differentiation such that the inner portion of the fetal zone atrophies and the zonae glomerulosa and fasciculata develop.286 Fetal zone remodeling in the human is an apoptotic process.287 It has generally been thought that the adult cortical zones develop from the persistent definitive zone. However, there is no evidence of adrenal cortical insufficiency during the neonatal period when the postnatal remodeling process is at its peak. It is more likely that the nascent adult cortical zones are present and functional before birth. Indeed, morphologic and functional studies have identified rudimentary zonae glomerulosa and fasciculata during late gestation.288 ACTH secreted from the fetal pituitary is the principal trophic regulator of the fetal adrenal cortex. However, ACTH may not be acting directly. During the last two-­thirds of gestation, the fetal zone grows rapidly and produces large amounts of steroids, even though circulating ACTH concentrations do not rise significantly. Soon after birth, the fetal zone rapidly involutes but exposure to ACTH continues, albeit at lower concentrations. Other factors, possibly specific to the intrauterine environment, appear to play a role in the regulation of fetal adrenal cortical growth and function. Substances produced by the placenta (e.g., CG) have been implicated, and peptide growth factors produced locally within the fetal adrenal appear to influence fetal adrenal cortical growth and function by mediating or modulating the trophic actions of ACTH.289,290 Evidence of fetal adrenal steroidogenesis is first seen at 6 to 8 weeks when the cells in the developing adrenal differentiate and acquire steroidogenic characteristics. At around week 12 of gestation, estriol concentrations in the maternal circulation rapidly increase (approximately 100-­fold). This increase coincides with the initiation of fetal zone enlargement and ACTH secretion by the fetal pituitary gland.282 Production of DHEA-­S by the fetal adrenal cortex continues for the remainder of the pregnancy and increases considerably during the second and third trimesters. By term, the human fetal adrenal produces around 200 mg DHEA-­S/ day. Thus, placental production of estriol directly reflects the steroidogenic activity of the fetal HPA axis. For this reason, maternal estriol previously was used as an endocrine marker to evaluate the status of the fetal HPA axis. The point at which the fetal adrenal cortex begins producing physiologically relevant amounts of cortisol has yet to be determined definitively. This question has been partially answered, however, by observations of infants with congenital adrenal hyperplasia (CAH). In CAH due to a deficiency of the 21-­hydroxylase (P450c21) enzyme, the fetal adrenals cannot synthesize cortisol.291

to loss of glucocorticoid-­negative feedback. Female infants with CAH often are born with urogenital sinus defects and virilization primarily caused by exposure of the urogenital sinus to excessive amounts of adrenal androgen early (∼week 8) in gestation. Because this condition derives from adrenal glucocorticoid deficiency, the observations imply that the adrenal produces enough cortisol in female fetuses before 10 weeks of gestation to regulate ACTH levels and to prevent overproduction of adrenal androgens that may masculinize the female urogenital sinus. Expression of key steroid metabolizing enzymes suggests that the human fetal adrenal cortex does not produce cortisol de novo from cholesterol until around week 30 of gestation.288 However, this does not preclude the possibility that cortisol is produced by using progesterone as a precursor early in gestation.292 Because the fate of pregnenolone metabolism is initially determined by the branch point steroidogenic enzymes P450c17 and 3β-­HSD, the steroidogenic potential of cells may be inferred by the pattern of expression of these two enzymes (Fig. 11.7). Expression of 3β-­HSD by the human fetal adrenal cortex is a critical step in the metabolism of pregnenolone because it confers on cells the ability to convert Δ5–3β-­ hydroxysteroids to Δ4–3 ketosteroids essential for mineralocorticoid and glucocorticoid production. Between 12 and 22 weeks of gestation, the human fetal adrenal cortex does not express 3β-­HSD. After 22 weeks, 3β-­HSD expression can be detected first in the definitive zone cells and later in gestation in the definitive and transitional zone cells. At no time in gestation is 3β-­HSD expression detected in the fetal zone. In contrast, expression of P450c17 is highly abundant in the transitional and fetal zones and is lacking in the definitive zone at all gestational ages.288,293 The persistent lack of 3β-­ HSD and expression of large amounts of P450c17 in the fetal zone is consistent with this cortical compartment producing only C19/Δ5 steroids, particularly DHEA. The lack of P450c17 in the definitive zone, and the eventual expression of 3β-­HSD in this compartment, is consistent with this zone producing mineralocorticoids late in gestation. Coexpression of 3β-­HSD and P450c17 in the transitional zone indicates that this zone has the capacity for cortisol production (Fig. 11.10). Steroidogenic enzymes downstream of P450c17 and 3β-­HSD also have been examined.294 Their localization and ontogeny are consistent with the concept that the definitive zone develops to form the zona glomerulosa, the transitional zone is analogous to the zona fasciculata, and the fetal zone is analogous to the zona reticularis. Mineralocorticoid production by the primate fetal adrenal cortex is low early in gestation but increases during the third trimester. At term, 80% of the aldosterone in human and rhesus monkey fetal blood appears to originate from the fetal adrenal.295 In 18-­to 21-­week human fetal adrenals, the mineralocorticoid metabolic pathway is localized to the definitive zone, but its activity is low and unresponsive to secretagogues.296,297 The angiotensin II receptors, AT1 and AT2, are present on human fetal adrenal cortical cells after 16 weeks.298 The AT2 receptor is localized mainly on definitive zone cells, whereas the AT1 receptor is detectable to a lesser extent in cells from both fetal and definitive zones. Thus, during the first and second trimesters, the ability of the human fetal adrenal cortex to synthesize mineralocorticoids is minimal even though the cells express AT receptors.

Fetal Pituitary-­Gonadal Axis The foundation for normal puberty and adult reproductive function is established during fetal life with the development of the fetal hypothalamic-­ pituitary-­ gonadal axis. Impairment of this system can lead to irreparable loss of germ cells and reproductive

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CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

Midgestation P450scc P450c17 P450c21 3β-HSD P450c11 Late gestation P450scc P450c17 P450c21 3β-HSD P450c11 Fig. 11.10 Schematic representation of the localization of expression of P450scc, 3β-­hydroxysteroid dehydrogenase (3βHSD), P450c17, P450c21, and P450c11 in the primate fetal adrenal cortex during midgestation and late gestation. The thickness of the line indicates the relative abundance of expression. Dashed line indicates a lack of expression. Note the lack of P450c17 expression in the definitive zone at all stages of gestation and the ontogenetic expression of 3βHSD only in the definitive and transitional zones late in gestation. (From Mesiano S, Jaffe RB. Developmental and functional biology of the primate fetal adrenal cortex. Endocr Rev. 1997;18:378–403)

Studies of anencephalic fetuses indicate that embryonic sexual differentiation and early gonadal development do not depend on fetal pituitary gonadotropes. In anencephalic male fetuses, there is a reduction in Leydig cell number, whereas the number of spermatogonia is normal.299 There is also a similar time of appearance of seminiferous tubules in anencephalic and normal fetuses. In female anencephalic fetuses, the ovaries appear to develop normally until 32 weeks. At term, ovaries of anencephalic fetuses are smaller than normal, central follicles are absent, and proliferation of oogonia with progression through meiosis until the development of primordial follicles is not initiated.299 Thus, pituitary gonadotropins are necessary for granulosa cell-­follicular proliferation near term; they are also needed to a lesser extent, via granulosa cell regulation, for oocyte survival. On the basis of studies in fetal rhesus monkeys, fetal pituitary gonadotropins appear necessary for granulosa cell proliferation and follicular fluid formation during the latter part of pregnancy.300 Fetal testes exhibit steroidogenic activity and synthesize testosterone de novo.301 Testosterone secretion by Leydig cells is necessary at different stages of pregnancy for adequate growth and differentiation of the male internal and external genitalia. In the fetal rhesus monkey in utero, testosterone produc tion can be elicited by CG injection and by stimulating the fetal

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pituitary-­testicular axis with GnRH.276,302 Unsurprisingly, aromatase activity is low in the fetal testis.303 In contrast to the male, ovarian steroid hormone production is not essential for female phenotypic development. Furthermore, meiosis in the ovary begins relatively early, but significant estrogen production does not occur until late in fetal life, although the capacity for aromatization exists by the eighth week.304 The preponderance of data indicates that although certain enzymatic activity can be demonstrated in the fetal ovary, it cannot form estrogen de novo and is probably steroidogenically quiescent through most of pregnancy. As with the fetal adrenal cortex, locally produced growth factors, acting in an autocrine-­paracrine manner, appear to play critical roles in controlling the growth and development of the fetal gonads.305 The development of the fetal testis is described further in Chapter 17 and that of the fetal ovary in Chapters 8 and 17.

Fetal Pituitary-­Thyroid Axis In the human fetus, the thyroid gland acquires its characteristic morphological appearance and the capacity to concentrate iodine and synthesize iodothyronines by 10 to 12 weeks of gestation. Coincidentally, hypothalamic TRH is detectable at 10 to 12 weeks when thyrotropes can be detected in the fetal pituitary and TSH can be found in the fetal pituitary and serum. Thyroxine (T4) has also been detected in the fetal circulation.306 Thyroid function remains in a basal state until midgestation. At this time, the secretory activity of the thyroid gland and serum T4 concentrations begin to increase. This rise is likely to be related to the establishment of continuity between the hypothalamic and pituitary portions of the portal vascular system. Pituitary and serum TSH concentrations begin to rise shortly before the rise in T4 levels. Maximal TSH concentrations are reached early in the third trimester and do not increase further until term. In the rhesus monkey, a TSH response to TRH administration is present early in the equivalent of the third trimester.307 In contrast to the adult, triiodothyronine (T3) administration to the fetus does not suppress the pituitary TSH response to TRH in primates.307 The human infant born after 26 to 28 weeks of gestation responds to exogenous administration of TRH with an increase of circulating TSH levels similar to that which occurs in adults.308 At term, TSH secretion by the human fetus can be inhibited by the administration of T4.308,309 In addition, the elevated cord T4 is associated with marked suppression of the neonatal TSH surge.309 This inhibitory effect of T4 is presumably mediated through pituitary conversion of T4 to T3. Human fetal serum T4 and free T4 levels increase progressively during the last trimester, although serum TSH levels do not.309 Serum T3 concentrations are usually not measurable in the human fetus until approximately the 30th week of gestation and rise gradually thereafter.309 The prenatal increase in serum T3 concentration occurs during several weeks and may be related to increased cortisol concentration.310 Immediately after birth (during the first 4–6 hours), circulating T3 levels increase still further to concentrations three to sixfold those occurring in utero.310,311 The level of reverse T3 (3, 3’, 5’-­triiodothyronine) is high (∼250 ng/dL) in the human fetus early in the third trimester and then decreases progressively until term and remains virtually unchanged during early neonatal life in term infants.309,311,312 An acute increase in pituitary TSH levels occurs when the term fetus is exposed to the extrauterine environment. This, in turn, stimulates thyroidal iodine uptake and evokes the release of thyroid hormones. The maximal TSH concentrations are attained 30 minutes after birth. Thereafter, there is a rapid decrease in serum TSH during the first day of extrauterine life and a slower decrease during the succeeding 2 days. Serum T4 and free T4

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levels reach a peak at 24 hours and then decrease slowly during the first weeks of life.313 The binding of iodothyronine and the maturation of thyroid hormone receptors have not been reported in the human fetus. In the rat, hepatic nuclear T3 receptors mature during the first few weeks after birth.314 T3 receptor capacity, and possibly affinity, increases during the first 3 to 4 weeks. In the brain (in contrast with the liver), T3 nuclear receptor binding develops early, is comparable with that in the adult brain, and increases still further in the first 3 days of neonatal life.315 However, there appears to be a discrepancy between T3 nuclear receptor binding and brain tissue responsiveness in both the neonate and the adult. Neither appears to respond to exogenous T3 administration with an increase in oxygen, α-­glycerophosphate dehydrogenase, or malic enzyme consumption.315 Of interest is the observation that thyroid hormones increase nerve growth factor concentrations in adult and newborn mouse brain316 and it has been postulated that nerve growth factor mediates the effects of thyroid hormones on brain development.315

Fetal Pituitary-­Growth Hormone Axis GH is produced and secreted by the human fetal pituitary from around 8 to 10 weeks of gestation.317 Levels of GH in the fetal circulation peak at around 6 nmol/L at midgestation, then gradually decrease for the remainder of gestation to around 1.5 nmol/L at term. The peak in GH at midgestation is thought to be due to unrestrained secretion because cultured pituitary cells from 9-­to 16-­week fetuses respond to GHRH with increased GH secretion and are far less responsive to inhibition by SS.318 Alternatively, negative feedback pathways for the regulation of GH secretion may develop later than GHRF stimulation. The physiological control of pituitary GH production and secretion is mature by term. The role of fetal pituitary GH is unclear. Anencephalic fetuses on average are of normal size and weight, suggesting that fetal growth is controlled by factors other than fetal pituitary GH. Placental GH and PL are likely candidates. Clearly, pituitary GH is required for postnatal growth and development, and therefore this system must be mature at birth. However, it likely plays a minimal role in the control of fetal somatic growth.

Fetal Pituitary-­Prolactin Axis Levels of prolactin in fetal circulation are very low during the first half of human pregnancy. The pituitary begins secreting prolactin at 25 to 30 weeks, gradually secreting more for the remainder of gestation, and reaches a peak at term.319 The level of prolactin expression by the fetal pituitary increases progressively from around 15 weeks. In anencephalic fetuses, pituitary prolactin content is within the normal range, indicating that its production by the pituitary is independent of hypothalamic control during most of gestation.319 Control of prolactin secretion by TRH and dopamine matures late in pregnancy and during the first few months after birth.317,319 In cultured pituitary cells obtained from midgestation of human fetuses, estrogen stimulates and dopamine inhibits prolactin secretion.317 Interestingly, the increase in fetal prolactin levels with ongoing gestation coincides with increases in estrogen levels, indicating that its production by the fetal pituitary is influenced by estrogen during most of pregnancy. The role of prolactin in fetal development remains uncertain.

FETAL MATURATION AND THE TIMING OF PARTURITION • The appropriate timing of birth is essential for the success of pregnancy.

• Birth timing is linked to fetal neuroendocrine development so that the fetus is born only when it is physiologically equipped to survive as a newborn. • The process and timing of parturition involve specific endocrine and paracrine signals between the fetal and maternal compartments that control the labor state (i.e., quiescent or in labor) of the uterus.

Fetal Organ Maturation and Preparation for Extrauterine Life At birth, the fetus is abruptly required to establish and maintain physiologic homeostasis independently of the placenta and in a markedly altered environment. Survival of the neonate is therefore dependent upon the functional maturation of organ systems during fetal life that will be essential for extrauterine life. Critical among these are organs that interface with the environment and extract resources (e.g., lungs, gut, and immune system) and those that maintain homeostasis (e.g., HPA axis, kidneys, liver, and pancreas). The coordination of fetal maturation with birth timing is therefore critical for neonatal survival and pregnancy success. Many studies in multiple species have demonstrated that cortisol promotes the functional maturation of fetal organ systems.320 Critical processes induced by cortisol include surfactant production by the fetal lungs; activity of enzyme systems in the fetal gut, retina, pancreas, thyroid gland, and brain; and deposition of glycogen in the fetal liver. In sheep, fetal organ maturation is induced by a prepartum surge in cortisol secretion by the fetal adrenals.321 The cortisol surge also triggers the onset of labor (discussed later in the chapter). Thus, in sheep, the fetal HPA axis, via cortisol, mediates a physiologic link between the timing of birth and fetal organ maturation. The effect of cortisol (and synthetic glucocorticoids) on the maturation of the fetal lungs is especially important. The inability to exchange gases due to pulmonary immaturity (respiratory distress syndrome) is the leading cause of neonatal morbidity and mortality among preterm infants. Synthetic glucocorticoids that readily cross the placenta are administered to women in preterm labor as a standard of care to accelerate the maturation of fetal organ systems, especially the lungs. This treatment significantly increases survival rates among preterm infants mainly by promoting lung maturation and decreasing the severity of respiratory distress syndrome. The extent to which cortisol from the human fetal adrenals regulates fetal organ maturation is uncertain. Experiments of nature suggest that the maturation of the human fetus during late gestation is independent of fetal adrenal cortisol production. Fetuses with CAH due to P450c21 deficiency produce markedly reduced levels of cortisol, yet these infants are usually born at term without any apparent signs of lung immaturity.322 This observation suggests that cortisol produced by the fetal adrenal gland is not essential for fetal organ maturation and that another source of glucocorticoid, possibly from the maternal adrenals, could contribute to the stimulation of fetal organ maturation at the end of human gestation. It is also possible that the maturation of the human fetus is not dependent upon cortisol alone; other factors may be involved. Although cortisol stimulates the maturation of fetal organ systems, it also can have adverse effects on fetal development. To protect the fetus from these negative effects, the human placenta prevents maternal cortisol from entering the fetal compartment throughout most of the human pregnancy by expressing the 11β-­ HSD-­2 enzyme that converts cortisol to the inactive cortisone.241 Thus, for most of pregnancy, the placenta forms a biochemical barrier to maternal cortisol. Late in human pregnancy (30–35 weeks), however, the placental barrier to maternal cortisol weakens. The evidence for this is that estriol levels in the maternal circulation during late pregnancy are inversely related to the circadian changes in circulating maternal cortisol levels.323 Thus,

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

when maternal cortisol goes up, estriol goes down. This implies that, late in gestation, some maternal cortisol crosses the placenta to the fetal compartment and suppresses ACTH production by the fetal pituitary gland that leads to decreased DHEA-­S production by the fetal adrenals and in turn decreased estriol production by the placenta. Increased transfer of maternal cortisol to the fetus may represent a backup mechanism to ensure fetal lung maturation. This may explain why fetuses with cortisol deficiency are born without overt signs of organ system immaturity. This level of redundancy is expected for a system that is mission-­ critical for the success of reproduction. The link between fetal adrenal cortisol and the timing of birth is not as stringent in humans as it is in sheep. Human parturition appears to be independent of fetal HPA activity, although a role for placental CRH modulation of fetal adrenal DHEA production may be involved (see below).

Process of Human Parturition The process of parturition involves: (1) transformation of the myometrium from a quiescent to a highly contractile and excitable state to produce the contractions of labor; (2) remodeling of the uterine cervix such that it softens and dilates to allow passage of the fetus; and (3) weakening and rupture of the fetal membranes. These temporally coordinated events can be divided into distinct phases (Fig. 11.11).324 Phase 0 (quiescence): For most of pregnancy, the uterus is in a state of relaxation and relatively insensitive to stimulatory uterotonins (factors that affect uterine contractions); for example, prostaglandins (PGs) and oxytocin (OT), and the uterine cervix remains closed and rigid. This period, referred to as Phase 0 or quiescence, is controlled by propregnancy relaxatory uterotonins such as β-­adrenergic agents, PGI2, relaxin, CRH, parathyroid hormone-­related peptide and nitric oxide, and uterotropins (factors that modulate uterine function and growth), especially progesterone. In general, factors that activate adenylate cyclase or guanylyl cyclases and increase intracellular cAMP and cGMP, respectively (e.g., via Gαs-­protein-­coupled receptors) promote myometrial relaxation. Phase 1 (transformation): As an essential prelude to active labor, the myometrium undergoes a phenotypic transformation, referred to as Phase 1, whereby it gains the capacity to contract forcibly and rhythmically and becomes more responsive to uterotonins. The awakening of the quiescent uterus is initiated by progesterone withdrawal and increased estrogenic effects leading to distinct biophysical changes in the myometrium and cervix. Birth

Progesterone

Progesterone withdrawal

Uterotonins (PGs, OT)

Uterotonins (OT)

Estrogen activation Myometrial contractile activity Phase 0 20-40 weeks

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The myometrium transformation involves the expression of a specific cohort of genes, collectively referred to as contraction-­ associated proteins (CAPs).325 Some important CAPs include connexins (e.g., connexin-­43), which form gap-­junctions between myometrial cells, allowing rapid electrical and chemical signaling between adjacent cells to permit coordinated contractions; ion channels (e.g., calcium channels), which determine the resting membrane potential and excitability; and uterotonin receptors (e.g., OT and PG receptors). Concurrently, the uterine cervix gradually softens and becomes more elastic due to increased expression of collagenase and metalloproteinase enzymes that remodel the cervix extracellular matrix. In addition, intrauterine production of PGE2 and PGF2α increases due to increased PTGS2 activity by the fetal membranes and decidua. Phase 2 (activation): The increased myometrial exposure to PGs and its augmented responsiveness to PGs and OT eventually initiate Phase 2, which is defined by the onset of active labor. During Phase 2, uterotonic drive induces coordinated rhythmic contractions that eventually become more forceful. At the peak of active labor, the myometrium is one of the strongest muscles (on a per weight basis) in the human body. As the cervix effaces and becomes more compliant it dilates completely in response to the contractions of labor that progressively move the fetus and placenta through the birth canal. Phase 3 (hemostasis and involution): Phase 3 begins after the placenta, fetal membranes, and decidua (endometrium of pregnancy) are expelled. Importantly, myometrial contraction is sustained (mainly in response to OT) during this time, which helps constrict the spiral arterioles to facilitate postpartum uterine hemostasis. During the following weeks, the uterus gradually reverts back to the menstrual state through a combination of myometrial cell apoptosis and atrophy and reformation of the cycling endometrium. In addition, the cervix remodels and reverts to a closed and rigid state.

Hormonal Control of Human Parturition Parturition is controlled by multiple hormones that affect the growth and contractility of the myometrium and the mechanical integrity of the uterine cervix and fetal membranes. Central to the process are progesterone and estrogens (mainly estradiol). For most of pregnancy, progesterone actively blocks parturition and promotes myometrial relaxation and cervical closure. Parturition is triggered when the progesterone block to labor is removed. Withdrawal of progesterone allows the uterus to transition to the laboring state to empty its contents. Estrogens, in contrast, oppose the actions of progesterone by stimulating biochemical and physical changes in the myometrium, cervix, and fetal membranes that promote labor. The labor status of the pregnant uterus is therefore thought to be determined by the balance between propregnancy and labor-­ blocking actions of progesterone and prolabor actions of estrogens. During most of pregnancy, the propregnancy actions of progesterone (i.e., the progesterone block to labor) prevail, and parturition is triggered by progesterone withdrawal.

Progesterone withdrawal Phase 1

Phase 2

Phase 3

1-2 weeks

Fig. 11.11 Phases of human parturition based on myometrial contractile activity and the principal regulatory uterotropins and uterotonins involved. (Adapted from Casey ML, Macdonald PC. Endocrine changes of pregnancy. In: Wilson JD, Foster DW, Kronenberg HM, Larsen PR, eds. Williams Textbook of Endocrinology. W.B. Saunders; 1998:1259.)

Numerous studies in multiple species demonstrate the pivotal role of progesterone withdrawal as a trigger for parturition. In all viviparous species studied so far, parturition is induced by any intervention that interferes with progesterone synthesis or action, and in most species, normal parturition at term is triggered by a prepartum decrease in circulating progesterone levels.326 The mechanism for progesterone withdrawal and how it is controlled differs between species. In some animals (e.g., rodents), the CL is the exclusive source of progesterone though pregnancy and parturition are caused by its regression leading to systemic

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progesterone withdrawal. In other species (e.g., ovine, bovine), progesterone is produced by the placenta and progesterone withdrawal at parturition is caused by hormonal signals from the fetus that decrease placental progesterone secretion. In sheep, this is induced by a surge in cortisol production by the fetal HPA axis 1 to 2 weeks before term.327 The fetal cortisol surge stimulates the expression of the P450c17 enzyme in the placenta which converts progesterone to androstenedione leading to systemic progesterone withdrawal.328 This also leads to a concomitant increase in placental estrogen production due to the increased availability of androstenedione for aromatization (see Fig. 11.7). The fall in circulating progesterone levels, coupled with a rise in circulating estrogens, transforms the uterus to the laboring state. As cortisol is the principal stimulator of fetal organ system maturation, the prepartum fetal cortisol surge ensures that parturition is coordinated with fetal maturation. Although cortisol promotes the functional maturation of fetal organ systems in the human, as it does in sheep, its production by the fetal HPA axis does not appear to be necessary for normal human parturition.322,329 As described above, parturition occurs normally at term in pregnancies where the fetus has CAH due to the inability to produce cortisol.322 Transfer of cortisol from the maternal to the fetal compartment may compensate for the lack of fetal adrenal cortisol in CAH pregnancies, but whether it plays a role in triggering parturition is unlikely since the administration of glucocorticoids to women in threatened preterm birth does not appear to advance the parturition process. A distinguishing feature of parturition in humans and some nonhuman primates is that circulating levels of progesterone and estrogens are high for most of pregnancy and during labor and delivery (see Fig. 11.9).330 Thus, in human parturition, the uterus transitions to the laboring state despite being exposed to high levels of progesterone, and for most of pregnancy, the uterus remains quiescent despite being exposed to high levels of estrogens. Nonetheless, labor in women can be induced at any stage of pregnancy by interventions (e.g., administration of mifepristone, a progesterone receptor antagonist) that inhibit progesterone action.331 One hypothesis to explain this dynamic is that human parturition involves a functional, rather than systemic, progesterone withdrawal, whereby uterine cells become refractory to progesterone actions and this is linked with functional estrogen activation, whereby uterine cells become sensitive to the stimulatory prolabor actions of estrogens. Based on this paradigm the physiologic control of human parturition involves hormonal interactions that control progesterone and estrogen responsiveness in uterine target cells. Progesterone responsiveness is primarily determined by the extent of expression and transcriptional activity of the nuclear progesterone receptors (PRs) that belong to the nuclear receptor family of ligand-­ activated nuclear transcription factors.332 Transcription of the human PR gene (PGR) is controlled by two promoters that produce two major PR isoforms, PR-­A and PR-­ B. PR-­A is a truncated (by 165 N-­terminal amino acids) form of PR-­B. Progesterone target tissues coexpress PR-­A and PR-­B in varying relative amounts depending on tissue type and pathological condition, and as such responsiveness to progesterone is dependent on the combined actions of PR-­A and PR-­B. Initial studies examining the transcriptional activities of PR-­A and PR-­ B using the canonical progesterone-­ responsive DNA element showed that upon exposure to progesterone, PR-­B has strong transcriptional activity, whereas PR-­A on its own has minimal transcriptional activity and, depending on its amount relative to PR-­B, represses the transcriptional activity of PR-­B.333 Those findings led to the concept that PR-­A and PR-­B comprise a dual system for the control of progesterone responsiveness, whereby progesterone responsiveness is inversely related to the cellular PR-­A/PR-­B ratio. This increased PR-A relative to PR-B in uterine cells could cause functional progesterone withdrawal.

Consistent with this hypothesis several studies found that PR-­A represses the transcriptional activity of PR-­B in human myometrial cells and that the onset of labor is associated with a significant increase in myometrial PR-­A.334–339 Other mechanisms for functional progesterone withdrawal also have been described. Progesterone responsiveness in uterine cells could be inhibited by increased expression of PR-­B corepressors, the interaction of PR with other transcription factors, and reduced binding of PRs to DNA.340–343 Studies also suggest that progesterone is catabolized to an inactive form that fails to bind to the PRs in myometrial cells thus causing the PRs to become unliganded.344–347 In vitro studies suggest that unliganded PR-­A stimulates the expression of CAP genes to augment myometrial contractility.347 It is also possible that specific micro RNAs affect PR signaling, leading to increased expression of CAP genes.345,348 Thus, current data indicate that functional progesterone withdrawal in human parturition is mediated by a variety of processes that abrogate the capacity for ligand-­activated PR-­B to exert genomic actions that promote uterine quiescence. Thus, as with fetal maturation in preparation for extrauterine life, redundancy in the triggers for parturition, that manifest as multiple pathways for progesterone withdrawal, would be expected since ending pregnancy is critical for the success of pregnancy and survival of the female.

Inflammation and progesterone withdrawal Multiple studies indicate that human parturition is an inflammatory process and that labor is associated with tissue-­level inflammation within the myometrium, cervix, and decidua.349–353 A significant proportion of preterm births are associated with intrauterine infection, clinically silent upper genital tract infection, and bacterial vaginosis.354–361 In rhesus macaques, inflammation in the gestational tissues precedes the onset of labor362,363 and administration of proinflammatory stimuli into the uterus initiates preterm labor.364–366 Thus, it is generally considered that inflammatory stress induces parturition. Progesterone exerts antiinflammatory activity and this is considered to be the principal mechanism by which it blocks labor.367–369 In myometrial cell cultures, progesterone via PR-­B inhibits the responsiveness to proinflammatory stimuli,370–372 and this effect is inhibited by PR-­A.369 Studies in immortalized human myometrial cell lines show that proinflammatory cytokines increase the abundance of PR-­A and its capacity to repress PR-­ B373 and that this is likely caused by site-­specific phosphorylation of PR-­A.374 The data suggest that inflammatory stimuli induce labor by activating PR-­A-­mediated functional progesterone withdrawal. It is proposed that for most of pregnancy, progesterone/ PR-­B antiinflammatory effects in uterine target cells dominate to block inflammation-­induced parturition and that an inflammatory load threshold exists above which PR-­A-­mediated functional progesterone withdrawal is induced (Fig. 11.12).374 The model posits that inflammatory stimuli from physiologic and pathologic sources gradually accumulate with advancing gestation toward the threshold for inducing PR-­A transrepressive activity. The ensuing inflammation leads to increased local production of PGs that induce labor by increasing myometrial contractility and softening the cervix (Fig. 11.13). This mechanism explains, at least in part, the increased incidence of preterm birth associated with infection/inflammation and fetal/maternal stress.

Progestin therapy for Preterm Birth Prevention The discovery of the natural progestation and labor-­blocking actions of progesterone in the 1930s logically led to its clinical use to prevent preterm birth. Even though it was known that progesterone levels in human pregnancy are high, it was reasoned that supplementing progesterone would promote uterine

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

Prolabor signal s Uterine distention Chorioamnionitis Fetal stress Maternal stress Abruption Other...

Progesterone

Progesterone PR-B

PR-B

PR-A

Labor

PR-A-P Cytokines PGs Tissue-level inflammation

Functional progesterone withdrawal

Quiescence Inflammatory load threshold

Inflammatory load Time

Time

TERM

TERM

Fig. 11.12 Theoretical model for the functional interaction between inflammatory load and progesterone responsiveness with advancing gestation. For most of pregnancy, progesterone acting via the PR-­B promotes uterine quiescence. With advancing gestation prolabor signals from multiple sources (listed in box) increase the inflammatory load on the pregnancy uterus until a threshold is reached above which protein kinase pathways are activated in myometrial cells that phosphorylate PRA, which activates repressive activity on PR-­B to cause functional progesterone withdrawal. According to this paradigm, the timing of parturition is determined by the trajectory of the inflammatory load curve and the set-­point for the inflammatory load threshold. Triggers for parturition Uterine stretch

Infection/ inflammation

Other

PGs are direct uterotonins PGs

Cytokines

PTGS2

Cytokines induce phosphorylation of PR-A that activates its repressive activity of PR-B

Cytokines augment PG production by myometrial cells via the NF B pathway

NF B

P

Estrogens augment contractility by stimulating CAP gene expression via ER ER

PR-A

PR-B

Estrogens

CAPs

PR-B inhibits NF B activity, CAP gene expression, and estrogen responsiveness

Contractile state

Progesterone Myometrial cells

Fig. 11.13 Theoretical model for how progesterone maintains uterine quiescence and how various physiological pro-­birth inputs initiate the human parturition cascade. In myometrial cells progesterone via PR-­B promotes relaxation by inhibiting responsiveness to pro-­inflammatory stimuli, inhibiting CAP gene expression, and inhibiting responsiveness to estrogens. As the pregnancy advances, the inflammatory load on the uterus increases. When an inflammatory load threshold is reached, cytokines activate a kinase that phosphorylates PR-­A which activates its repressive effect on PR-­B to cause functional progesterone withdrawal. Removal of the progesterone/PR-­B block causes tissue-­ level inflammation and increased production of local PGs that promote myometrial contraction and cervical softening.

relaxation and prevent preterm birth. Trials of high doses of progesterone (as a bolus intravenously or into the amniotic fluid) to decrease the frequency of spontaneous contractions showed that this approach does not inhibit established labor and consequently progesterone is not used for tocolysis. Several studies examined the use of prophylactic therapy with a synthetic caproate ester of 17α-­hydroxyprogesterone (17HPC) and showed that it may reduce the risk for preterm birth.375–377 Larger clinical trials of 17HPC and progesterone were subsequently conducted and it was found that prophylactic treatment beginning at around midg estation with micronized progesterone administered vaginally

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or 17HPC given weekly via intramuscular injections379 decreased the incidence of preterm birth and improved neonatal outcome. This effect, however, was limited to women with an increased risk for preterm birth. 17HPC was effective only in women with a history of preterm birth and micronized progesterone was effective only in women with a short cervix assessed by ultrasound at midgestation.380–383 To confirm this efficacy, larger trials were conducted for each treatment. Those studies found that neither treatment decreased the risk for preterm birth.384,385 The use of progestin prophylaxis for preterm birth prevention remains controversial.

Estrogen activation Estrogen signaling in uterine cells is considered to oppose the labor-­ blocking actions of progesterone and in most species, maternal estrogen (mainly estradiol) levels rise prior to delivery. In humans, this rise begins at midgestation and increases gradually during the remainder of pregnancy (previously discussed in the chapter). The role of placental estrogens in the endocrine control of human parturition, however, is equivocal because congenital abnormalities affecting estrogen production by the fetal-­ placental unit, such as anencephaly, congenital adrenal lipoid hyperplasia, placental aromatase deficiency, and placenta sulfatase deficiency, do not affect the timing of birth. It appears that relatively little estrogen is required for normal human pregnancy and parturition and elevated levels do not cause preterm birth. Thus, estrogen actions in human parturition are not controlled by levels in the maternal circulation. Instead, estrogen responsiveness, in this case increased responsiveness to estrogens (i.e., functional estrogen activation), is likely to be more important. Functional estrogen activation at parturition could be mediated by increased expression of estrogen receptor-­α (ERα) in uterine in target cells. Expression of ERα in term myometrium increases significantly in association with the onset of labor and correlates with increased expression of mRNA encoding connexin-­43, a key estrogen-­responsive CAP gene.334 Numerous studies in a variety of species have demonstrated a functional interaction between the ER and PR systems such that progesterone via PR-­B decreases uterine estrogen responsiveness by decreasing expression of ESR1 (encodes ERα), and estrogen via ERα increases uterine progesterone responsiveness by increasing expression of PGR (encodes PR-­A and PR-­B). Levels of ERα expression in quiescent (i.e., before labor) term human myometrium correlate positively with the PR-­A/PR-­B mRNA ratio,334 suggesting that myometrial cell expression of ERα increases as progesterone responsiveness decreases. Studies in the rhesus monkey have shown that inhibition of progesterone action with the PR antagonist, mifepristone (also known as RU486), increases ERα expression in the pregnancy myometrium.386 Thus, progesterone may decrease uterine estrogen responsiveness by inhibiting myometrial ESR1 expression. This would explain why the myometrium is refractory to the prolabor actions of estrogens for most of pregnancy. It also explains why functional withdrawal of progesterone/PR-­B signaling (e.g., by increased transrepressive activity of PR-­A) initiates the full parturition cascade.

Placental CRH A unique feature of primate pregnancy is that the placenta produces CRH.387 Studies of maternal CRH levels during human pregnancy suggest that placental CRH plays a role in the physiology of parturition. The trajectory of maternal CRH levels at midgestation is predictive of whether parturition will occur at term, preterm, or postterm (Fig. 11.14).106 Interestingly, the exponential rise in maternal plasma CRH concentrations with advancing pregnancy is associated with a concomitant fall in the binding protein in late pregnancy

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Maternal Plasma CRH (pmol/L)

100

CRH

Preterm Term Postterm

+

75

ACTH +

Uterus

CRH

50

Oxytocin receptors + Gap junctions Prostaglandins

+ +

Estrogens

Fetal hypothalamus

Fetal pituitary

+++ Cortisol

+++ +++ DHEA-S

+ Fetal lung

Placenta Fetal adrenal

25

0 15

20

25

30

Gestation (weeks) Fig. 11.14 Schematic representation of the association between changes in maternal plasma CRH levels between 15 and 30 weeks gestation and the timing of birth. (From McLean M, Bisits A, Davies J, et al. A placental clock controlling the length of human pregnancy. Nature Med. 1995;1:460.)

(see Fig. 11.5).106 The implication of these reciprocal concentration curves is that there is a rapid increase in circulating levels of bioavailable CRH concurrent with the onset of parturition. CRH may be involved directly in the regulation of human parturition by modulating myometrial contractility. Receptors for CRH have been identified in the human myometrium and fetal membranes.388–391 In vitro studies have shown that CRH stimulates the release of PGs from human decidua and amnion,392,393 and augments the action of OT and PGF2α on myometrial contractility.394–396 During most of pregnancy, CRH increases adenylate cyclase activity in the quiescent myometrium, leading to an increase in intracellular cAMP, which promotes relaxation. In contrast, at around the time of parturition the capacity for CRH to increase cAMP decreases, especially in laboring myometrium.397 Whether changes in CRH receptor signaling contribute to the onset of labor remains unknown. The extrahypothalamic source of CRH may have profound effects on the activity of the fetal-­placental unit. A positive feedback endocrine loop may develop between placental CRH and the fetal adrenal cortex as a consequence of the direct action of CRH on the fetal adrenal cortex and the stimulation of placental CRH expression by cortisol. In addition, CRH may increase ACTH responsiveness in fetal adrenal cortical cells (especially cells in the transitional zone) by increasing ACTH receptor expression.265 As a result, fetal adrenal cortisol and placental estrogen synthesis may increase through a positive feedback loop involving placental CRH (Fig. 11.15).

Prostaglandins PGs produced by intrauterine tissues are critical regulators of myometrial contractility and cervical softening (see Chapter 4).398–400 Production of PGE2 and PGF2α by intrauterine tissues (particularly the amnion, chorion, decidua, and myometrium) increases late in gestation and in association with the onset of labor, and administration of PGE2 or PGF2α at any stage of pregnancy induces uterine contractions and cervical ripening and causes labor and delivery.401,402 Inhibition of PG biosynthesis with aspirin, indomethacin, or specific PTGS2 inhibitors suppresses labor and prolongs gestation.403,404 The regulation of intrauterine PG production is considered to be a pivotal event in human parturition. The rate-limiting step in PG biosynthesis is catalyzed by two isozymes, PTGS1 and

Fig. 11.15 Schematic model of how placental corticotropin-­releasing hormone (CRH) modulates activity of the fetal-­placental unit. An increase in bioactive placental CRH at term has effects on both the fetus and the mother. In the fetus, CRH stimulates the fetal adrenal directly, to stimulate production of DHEA-­S, which is converted by the placenta to estrogen, which in turn drives expression of CAPs necessary for uterine contractions and parturition. The placental CRH may also act on the fetal adrenal cortex by up-­regulating ACTH receptor expression and increasing responsiveness to ACTH, which evokes fetal adrenal cortisol production from the definitive zone and DHEA-­S production from the fetal zone. A feed-­forward loop is thus started as fetal adrenal cortisol production, which in turn stimulates CRH production by the placenta. (From Smith R, Mesiano S, Chan EC, Brown S, Jaffe RB. Corticotropin-­releasing hormone directly and preferentially stimulates dehydroepiandrosterone sulfate secretion by human fetal adrenal cortical cells. J Clin Endocrinol Metab. 1998;83:2916–2920.)

PTGS2 (also known as cyclooxygenase-­1 and cyclooxygenase-­2) (see Chapter 4). PTGS1 is constitutively expressed in most tissue types, whereas PTGS2 expression is inducible, increasing in response to proinflammatory stimuli. PGs are metabolized by the prostaglandin dehydrogenase (PGDH) enzyme, which irreversibly converts PGE2 and PGF2α into inactive forms. Intrauterine biosynthesis of PGs occurs mainly in the amnion and, to a lesser extent, in the chorion. Both tissues express PTGS2, which increases in association with the onset of term and preterm labor. PGs also are produced in the decidua and in the myometrium, and in both sites, production increases at the time of labor. In the fetal membranes, PTGS2 expression increases during the third trimester and several weeks before the onset of labor.405 Exposure of the myometrium to fetal membrane-­derived PGs is dependent on the balance between PTGS2 and PGDH activities in the chorion. PGs produced by the amnion accumulate in amniotic fluid and do not necessarily gain access to target receptors on myometrial cells because the chorion, which lies between the amnion and the maternal tissues of the uterus, expresses high levels of PGDH.406 Thus, the chorion may act as a barrier throughout most of pregnancy, preventing amnion-­derived PGs from activating the myometrium (Fig. 11.16). Several studies have indicated that propregnancy factors such as progesterone prevent myometrial exposure to PGs by stimulating the expression of PGDH and inhibiting PTGS2. In contrast, cortisol, CRH, and several immune cytokines may inhibit PGDH and stimulate PTGS2, leading to a net increase in active PGs accessing the myometrium.398 Thus, the balance of PG synthesis and metabolizing activities in the fetal membranes may be pivotal in the control of uterine contractility. The pleiotropic actions of PGs in diverse tissue types can be explained by the existence of multiple receptor species linked to different intracellular signaling pathways.407 The regulation and localization of specific PG receptors in the myometrium and cervix are thought to be an important component in the hormonal control of pregnancy and parturition. During pregnancy, the human myometrium expresses receptors for PGE2, PGF2α,

Before parturition

CHAPTER 11  Endocrinology of Human Pregnancy and Fetal-Placental Neuroendocrine Development

PGHS PGDH

PGE2

Parturition

Amnion

Chorion

Decidua

Myometrium

PGF2α PGE2 PGE2

Amnion

PGF2α Chorion

Decidua

PGF2α PGE2

Myometrium

Fig. 11.16 Changes in prostaglandin endoperoxide H synthase (PGHS) and prostaglandin dehydrogenase (PGDH) activities in the fetal membranes decidua and myometrium associated with the onset of human labor. Parturition is associated with a marked increase in PGHS activity in all tissues. PDGH is predominantly expressed by the chorion and effectively blocks amnion prostaglandins (PGs) from accessing the myometrium. Parturition is associated with decreased PGDH activity in the chorion which could allow more active PG to reach the myometrium.

PGI2, and thromboxane.408 PGE2 interacts with four subtypes of PGE2 receptors (EPs): EP1, EP2, EP3, and EP4. In addition, eight splice variants of the EP3 receptor have been identified.409 With regard to myometrial contraction, the diversity of PGE2 receptor types may have important ramifications. Interaction of PGE2 with EP1 and EP3 increases intracellular calcium and decreases cAMP, leading to contraction. In contrast, the interaction of PGE2 with EP2 and EP4 activates adenylate cyclase, leading to increased cAMP and relaxation. Thus, PGE2 can cause uterine contraction or relaxation depending on the type of EPs expressed. In contrast, PGF2α interacts with a single receptor (FP) that increases intracellular calcium leading to contraction. PGI2 binds IP that is linked to adenylate cyclase, leading to increased intracellular cAMP and relaxation; although it may also increase CAP gene expression.399 Thromboxane binds to the thromboxane receptor (TP) and increases calcium and augments contractility. Studies of EP and FP expression in the term human uterus have shown that EP2 expression is greater than FP before the onset of labor and decreases with increasing gestation and that expression of the FP receptor is low before labor and increases significantly in association with the onset of labor.410 These findings suggest that myometrial relaxation through most of pregnancy is promoted, at least in part, by EP2, whereas the onset of labor involves an increase in myometrial FP, allowing PGF2α to stimulate contraction. Oxytocin OT, a 9-­amino acid peptide secreted by the posterior pituitary, is produced in some peripheral tissues and is one of the most potent stimulators of uterine contraction. It is routinely administered to promote labor and to induce tonic uterine contraction to decrease the risk of postpartum hemorrhage. OT is not considered to be involved in the initiation of labor because its

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circulating levels do not increase until the expulsive phase of labor.411 Inhibition of OT action by specific synthetic antagonists disrupts the normal pattern of labor and decreases uterine contractility in women with threatened preterm birth.412,413 Vaginal distention is a principal stimulus for the release of pituitary OT in human labor, via a neurologic feedback mechanism known as the Ferguson reflex. OT is produced by the amnion, chorion, and decidua and may have a local action on the adjacent myometrium.414 OT action appears to be regulated at the level of target tissue responsiveness by changes in OT receptor (OXTR) expression. In the human, myometrial and decidual OXTR content and expression increase gradually toward the end of pregnancy and then rise significantly in association with the onset of labor.415 Expression of the OXTR by myometrial cells is upregulated by estrogen.416 Thus, induction of OT responsiveness by increased OXTR expression may be the principal mechanism underlying the uterotonic actions of locally produced and pituitary OT in human pregnancy. After birth, pituitary oxytocin contributes to uterine involution and especially the tonic contraction of the myometrium needed for postpartum hemostasis. OT also affects the preparation of the breast for lactation (see Chapters 3 and 12).

Fetal Lung Maturation Studies in rodents have indicated that the fetal lungs contribute to the parturition process.417 In this species, surfactant protein -­A (SP-­A), a critical lung surfactant protein produced by the fetal lung epithelium, is secreted into the amniotic fluid where it activates fetal macrophages and causes them to migrate to the myometrium. In the myometrium, the increased production of cytokines by the fetal macrophages induces NFκB activation and tissue-­level inflammation leading to functional progesterone withdrawal. This interaction links fetal lung maturation with birth timing. Evidence for a similar system is not apparent in the human and the trafficking of fetal macrophages in the myometrium in humans and mice differs substantially. However, some studies suggest that SP-­ A interacts directly with myometrial cells via a specific receptor to increase prostaglandin production and contractile capacity.418 Thus, SP-­A produced by the human fetal lungs may directly affect myometrial contractility and the capacity for the development of an inflammatory response in the gestational tissues. This may impact the sensitivity of the gestational tissues to inflammation-­induced parturition. Interestingly, the fetal membranes express SP-­A in response to glucocorticoids and SP-­ A induces prostaglandin synthesis by chorionic trophoblasts.419

UTERINE STRETCH Uterine stretch has been proposed as a signal for the induction of parturition. Gestation is shorter in twin pregnancies,420,421 presumably due to the increased distention of the uterine wall. Studies in rats have shown that distention of a nonpregnant uterine horn induces changes in CAP gene expression similar to those in the pregnant horn and that this is inhibited by progesterone.422–424 The stimulatory effect of stretch on CAP gene expression was also observed in cultured human myometrial cells.425–429 Stretch of cultured uterine myocytes also increased expression and release of IL-­8, a potent proinflammatory chemokine that promotes the infiltration of activated macrophages and neutrophils.426 The human uterus is subjected to a significant amount of distention, especially late in the third trimester. A possible scenario is that a threshold of uterine stretch exists above which the induction of IL-­8 expression leads to a tissue-­level inflammatory state that induces PR-A-mediated functional progesterone

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EVOLUTIONARY PERSPECTIVE • The extant physiology of pregnancy is derived from the natural selection of traits to maximize reproductive efficiency and fitness. • The hormonal control of human pregnancy and birth timing was influenced by selective pressure of encephalization.    The reproductive system is at the leading edge of natural selection and as such traits affecting the success of pregnancy were likely subjected to strong selective pressures. This would explain the remarkable variation between species around the common theme of viviparity. In this context, unique characteristics of human pregnancy derive from traits unique to the Hominid lineage. One of the most important Hominid traits in this regard is encephalization: increased complexity and relative size of the brain. Encephalization conferred significant advantages for Hominid fitness and as such, natural selection likely favored pregnancy traits that promote encephalization. However, encephalization also has reproductive costs associated with the risk of birthing a large-­headed fetus, referred to as the obstetric dilemma.430 Obstetric difficulties caused by cephalo-­pelvic disproportion at birth are predominately a human problem. In addition, the cost of provisioning the fetus with sufficient energy to support rapid brain growth, referred to as the energetics of gestation and growth hypothesis, also confers metabolic and energetic constraints on pregnancy.431 One solution to these problems was to select for parturition trigger mechanisms that cause birth to occur before fetal head circumference exceeds the pelvic canal capacity and before fetal energy demands exceed the mother’s energy supply capacity. Analyses of the ratio of neonatal brain size to body size and maturity of the neonate across extant primate species suggest that human gestation is relatively short among extant primates.432,433 The human neonate is severely altricial (relatively immobile, unable to obtain food independently, and must have adult care), big-­headed, and has significant head/brain growth in the first 6 to 12 months after birth. Thus, it appears that natural selection favored a parturition trigger mechanism(s) that shortened gestation. This could explain the unique characteristic of human parturition such as lack of systemic progesterone withdrawal and inflammation/stress-­induced functional progesterone withdrawal. The same reasoning can be applied to maternal adaptations for pregnancy that are required to provide sufficient energy to facilitate the rapid growth of the fetal brain. This could have been achieved by the evolution of deep placentation and invasion of the uterine spiral arteries to maximize access to maternal blood, and the secretion of placental hormones such as PL and PGH that induce maternal insulin resistance to increase the supply of glucose for the fetus. Thus, an evolutionary framing for the hormonal control of human pregnancy, parturition, and neuroendocrine development puts into perspective extant physiology and adverse pregnancy conditions. Natural selection operates at the population level to reconcile the benefits and cost of traits to reproductive fitness over multiple generations. Extant

pathophysiology of pregnancy (e.g., preterm birth, preeclampsia) most of which are unique to humans, that occur at the individual level, may arise from inefficiencies and defects in hormonal systems that were selected based on net effects at the population level and over multiple generations rather than the individual pregnancy. This concept expands our perspective of pregnancy complications such as preeclampsia, fetal growth restriction, gestational diabetes, and preterm birth, and complex processes such as the plasticity of fetal neuroendocrine development in response to environmental cues. Net benefit over the long term may increase cost to the individual (e.g., increased risk for preterm birth, gestational diabetes) over the short term.434 Thus, the hormonal control of human pregnancy and parturition and the plasticity of fetal neuroendocrine development were shaped by the unique natural history of human evolution. TOP REFERENCES

Burton GJ, Fowden AL, Thornburg KL. Placental origins of chronic disease. Physiol Rev. 2016;96:1509–1565. Delorme-­Axford E, Donker RB, et al. Human placental trophoblasts confer viral resistance to recipient cells. Proc Natl Acad Sci U S A. 2013;110:12048–12053. Evans NP, Bellingham M, Robinson JE. Prenatal programming of neuroendocrine reproductive function. Theriogenology. 2016;86:340–348. Fowden AL, Forhead AJ, Sferruzzi-­ Perri AN, Burton GJ, Vaughan OR. Review: Endocrine regulation of placental phenotype. Placenta. 2015;36(suppl 1):S50–59. Gellersen B, Brosens JJ. Cyclic decidualization of the human endometrium in reproductive health and failure. Endocr Rev. 2014;35:851–905. Haig D. Genetic conflicts in human pregnancy. Quart Rev Biol. 1993;68:495. Mesiano S, Jaffe RB. Role of growth factors in the developmental regulation of the human fetal adrenal cortex. Steroids. 1997;62:62–72. Mitchell MD, Peiris HN, Kobayashi M, et al. Placental exosomes in normal and complicated pregnancy. Am J Obstet Gynecol. 2015;213:S173–181. Nadeem L, Shynlova O, Matysiak-­Zablocki E, Mesiano S, Dong X, Lye S. Molecular evidence of functional progesterone withdrawal in human myometrium. Nat Commun. 2016;7:11565. Norman JE, Marlow N, Messow CM, et al. Vaginal progesterone prophylaxis for preterm birth (the OPPTIMUM study): a multicentre, randomised, double-­blind trial. Lancet. 2016;387:2106–2116. Rightmire GP. Brain size and encephalization in early to Mid-­Pleistocene Homo. Am J Phys Anthropol. 2004;124:109–123. Roberts JM, Hubel CA. The two stage model of preeclampsia: variations on the theme. Placenta. 2009;30(suppl A):S32–37. Romero R, Nicolaides KH, Conde-­Agudelo A, et al. Vaginal progesterone decreases preterm birth